FUNCTIONAL MEMBRANES FOR LIMITED PROTEIN DIGESTION AND
PHOSPHOPEPTIDE ENRICHMENT PRIOR TO MASS SPECTROMETRY
By
Yujing Tan

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
DOCTOR OF PHILOSOPHY
Chemistry
2012

ABSTRACT

FUNCTIONAL MEMBRANES FOR LIMITED PROTEIN DIGESTION AND
PHOSPHOPEPTIDE ENRICHMENT PRIOR TO MASS SPECTROMETRY
By
Yujing Tan
Mass spectrometry (MS) is the leading technique for rapid identification of proteins and
characterization of their posttranslational modifications. However, as with most analyses, the
success of MS methods depends critically on sample purification and pretreatment. Microporous
membranes are attractive as a platform for enrichment and modification of proteins and peptides
because convection and short radial diffusion distances in pores provide rapid analyte mass
transport to binding or catalytic sites. Additionally, variation of the transmembrane flow rate can
control the extent of reactions such as digestion. This research exploits facile layer-by-layer
adsorption to incorporate nanoparticles or enzymes in micropores and create functional
membranes. As tools for sample preparation prior to MS analysis, TiO2-modified membranes
enrich phosphopeptides and protease-containing membranes afford control over protein digestion.
Posttranslational protein phosphorylation is one of the most important mechanisms for
creating protein diversity in eukaryotic cells. However due to the relatively low abundance of
most phosphopeptides in proteolytic digests, phosphopeptide enrichment is a prerequisite step for
effective and efficient MS analysis of phosphorylation. Alternating adsorption of poly(styrene
sulfonate) (PSS) and TiO2 nanoparticles in porous nylon substrates creates TiO2-modified
membranes for rapid enriching of phosphopeptides from small-volume (10’s of L) samples.
These TiO2-modified membranes bind 540 nmols of phosphopeptides in a 22-mm diameter disc
and selectively isolate the phosphopeptides from -casein, -casein and ovalbumin digests even

in a 100-fold excess of non-phosphorylated BSA digests if 2 % TFA is the loading and washing
buffer. The combination of membrane enrichment and tandem MS reveals seven phosphorylation
sites from in vivo phosphorylated tau protein, which is associated with Alzheimer’s disease, and
identification of such sites might aid the design of therapeutic drugs that inhibit phosphorylation.
Using a similar approach to membrane modification, sequential adsorption of PSS and
proteases in porous nylon substrates yields enzymatic membrane reactors for limited protein
3

digestion. Although a high local enzyme density (~30 mg/cm ) and small pore diameters in the
membrane lead to digestion in < 1 s, the low membrane thickness (170 m) affords control over
residence times at the ms level to limit digestion. Most importantly, pepsin-containing
membranes afford control of peptides sizes by altering the flow rate. Apomyoglobin digestion
demonstrates that peptide lengths increase as the residence time in the membrane decreases.
Under denaturing conditions, limited membrane digestion of bovine serum albumin and
subsequent ESI-Orbitrap MS analysis reveal large peptides (3-10 kD) that increase the sequence
coverage from 53 % (2-s digestion) to 82 % (0.05-s digestion). Moreover, electron transfer
dissociation tandem MS on a large myoglobin proteolytic peptide (8 kD) provides a resolution of
1-2 amino acids, which suggests that this protease-modified membrane might find applications in
hydrogen-deuterium exchange or middle-down proteomics studies. In addition, using trypsinand -chymotrypsin-modified membranes, a limited proteolysis study of a large Arabidopsis
GTPase, ROOT HAIR DEFECTIVE 3, shows suitable probing for labile regions near the Cterminus to suggest what protein reconstruction might make RHD3 more suitable for
crystallization. These studies clearly demonstrate the potential of functional membranes for
rapid and controlled purification or pretreatment prior to MS analysis.

I dedicate this dissertation to my parents, Guisheng Tan and Linghua Wu,
for their love and support.

iv

ACKNOWLEDGEMENT
First I would like to thank my advisor Prof. Merlin Bruening for his guidance and support
over the past five years. More than an advisor, he is a mentor, collaborator and friend. He has
taught me to be a scientist and to seriously do things the right way. His attitude and professional
conduct inspired me to pursue the honor of this degree. Despite numerous failures and
disappointments along the way, his patience and trust encouraged me to continue and eventually
make it to the finish line. Though seemingly infrequent, the moments we achieved exciting
results and success together helped me work through the hard times.
I want to give special thanks to my second reader, Prof. Gavin Reid, who served as the
advisor in the mass spectrometry portion of this work. His knowledge, enthusiasm and exactness
prompted my progress in mass spectrometry step by step. I am grateful for all his helpful
suggestions and input on my work in this dissertation. I also wish to thank Prof. Gary Blanchard
and Prof. Dana Spence for being my committee members and for their suggestions on this
project.
I also would like to thank Profs. Min-Hao Kuo, Michael Garavito, Federica Brandizzi,
Jetze Tepe, Narayanan Parameswaran and Daniel Jones for collaborations and kind help on this
dissertation work. Similarly, I would like to acknowledge Drs. Yi Zheng, Dexin Sui, Xiao Zhou,
Amanda Palumbo, Teri Lansdell, Li Cui , Jinlan Dong and Wen Qin for their help and
contributions to this dissertation.
I am also very grateful to all my fellow group members, those who graduated before me,
who are fighting hard now and those who just started, for their sincere support and help.
Especially, Drs. Lu Ouyang, David Dotzauer and Weihan Wang helped me in the early stages
and brought me up to speed with my lab-work. As a longtime friend and roommate, Weihan is
v

like a big brother to me. His hard work and interest in science have always set a good example
for me.
Lastly I wish to express my deepest appreciation to my family and friends for their love
and support. As the only child, I am sorry for not being home for every Chinese traditional
holiday and family reunion in the past five years. Their understanding and support sustained me
so I could accomplish my goal: being a doctor. Thanks to the internet, we could “see” and talk
with each other weekly. For the friends in both China and the U.S., whenever outside school, I
appreciate that they stayed with me through the good times and put up with my complaints and
mood in the bad times.
Because of the people and stories involved, writing this acknowledgement is like
traveling back through the days I spent chasing this degree. Without the help and support from
the people mentioned and countless more, I could not finish this dissertation. Finally, allow me
to thank them all again with my sincerest gratitude.

vi

TABLE OF CONTENT
Page
LIST OF TABLES..................................................................................xi
LIST OF FIGURES.............................................................................. xii
LIST OF SCHEMES ............................................................................xvi
LIST OF ABBREVIATIONS ............................................................ xvii
Chapter 1. Introduction .......................................................................... 1
1.1 Mass spectrometry for proteomics ..................................................................1
1.1.1 Ionization techniques..................................................................................2
1.1.2 Mass analyzers ...........................................................................................5
1.1.3 Proteomics approaches ...............................................................................7
1.2 Phosphorylation: a key to protein function ...................................................12
1.3 Phosphoproteomics: detection of protein phosphorylation, enrichment
techniques and analysis strategy ...........................................................................15
1.3.1 Detection of protein phosphorylation ......................................................15
1.3.2 Enrichment of phosphopeptides ...............................................................17
1.3.3 MS Analysis Strategy ...............................................................................24
1.4 Immobilized enzyme reactors ........................................................................27
1.4.1 Immobilization methods...........................................................................28
1.4.2 Immobilization supports...........................................................................32
1.4.3 Immobilized trypsin and pepsin, reactors for protein digestion ..............34
1.5 Hydrogen-deuterium exchange and limited proteolysis for MS ...................37
1.5.1 HDX-MS using the top-down approach ..................................................39
1.5.2 HDX-MS using the bottom-up approach .................................................39
1.5.3 HDX-MS using multiple acidic proteases ...............................................41
1.5.4 Limited proteolysis for protein structure elucidation...............................41
1.6 Outline of Dissertation ...................................................................................43
1.7 References......................................................................................................45

Chapter 2. Phosphopeptide Enrichment with TiO2-Modified
Membranes ............................................................................................. 60
2.1 Introduction....................................................................................................60
vii

2.2 Experimental ..................................................................................................63
2.2.1 Materials and solutions ............................................................................63
2.2.2 Membranes and holders ...........................................................................64
2.2.3 Synthesis of titania nanoparticles .............................................................64
2.2.4 Membrane modification ...........................................................................64
2.2.5 Breakthrough curves and determination of binding capacity ..................65
2.2.6 Determination of phosphopeptide recovery using labeled peptides ........66
2.2.7 Protein digestion .......................................................................................66
2.2.8 Phophopeptide enrichment from protein digests .....................................67
2.2.9 Mass spectrometry and data analysis .......................................................67
2.3 Results and discussion ...................................................................................68
2.3.1 Characterization of TiO2-modified nylon membranes .............................68
2.3.2 Phosphopeptide binding capacity .............................................................71
2.3.3 Phosphopeptide elution ............................................................................73
2.3.4 Phosphopeptide recovery .........................................................................75
2.3.5 Phosphopeptide enrichment from a mixture of -casein, -casein, and
ovalbumin digests ..............................................................................................79
2.4 Conclusions....................................................................................................84
2.5 Acknowledgement .........................................................................................84
2.6 References......................................................................................................86

Chapter 3. Investigation of Phosphorylation in Tau Protein and the
Nuclear Factor Kappa B (NF-B) Pathway using Phosphopeptide
Enrichment withTiO2-modified Membranes ..................................... 90
3.1 Introduction....................................................................................................90
3.2 Experimental ..................................................................................................93
3.2.1 Materials and solutions, membranes and holders, membrane modification
and protein digestion ..........................................................................................93
3.2.2 Purification of p-tau .................................................................................93
3.2.3 Enrichment of phosphopeptides from digested p-tau ..............................94
3.2.4 Immunoprecipitation of p65 from THP-1 cells........................................94
3.2.5 Enrichment of on-bead digests of immunoprecipitated p65 and associated
proteins ...............................................................................................................95
3.2.6 In vitro phosphorylation of IB by GRK5 and IKK ...........................95
3.2.7 Mass Spectrometry and Data Analysis ....................................................96
3.3 Results and discussion ...................................................................................98
3.3.1 Enrichment of p-tau digests .....................................................................98
3.3.2 Phosphopeptide enrichment from on-bead digests of immunoprecipitated
p65 and associated proteins .............................................................................109
viii

3.3.3 Analysis of in vitro phosphorylation of IB by GRK5 .......................115
3.4 Conclusions..................................................................................................122
3.5 Acknowledgement .......................................................................................123
3.6 References....................................................................................................125

Chapter 4. Limited Proteolysis Via msec-Digestions in ProteaseModified Membranes .......................................................................... 130
4.1 Introduction..................................................................................................130
4.2 Experimental ................................................................................................133
4.2.1 Materials .................................................................................................133
4.2.2 Membrane and holder.............................................................................133
4.2.3 Modification of membranes with proteases ...........................................134
4.2.4 Controlled apomyoglobin digestion with a pepsin-modified membrane
134
4.2.5 Controlled BSA digestion with a pepsin-modified membrane ..............135
4.2.6 Mass spectrometry and data analysis .....................................................136
4.3 Results and discussion .................................................................................137
4.3.1 Membrane modification with protease ..................................................137
4.3.2 Apomyoglobin digestion with different membrane residence times .....138
4.3.3 ESI-Orbitrap ETD-MS/MS analysis of a large apomyoglobin peptide .144
4.3.4 In-membrane BSA digestion under denaturing conditions ....................147
4.4 Conclusions..................................................................................................152
4.5 Acknowledgement .......................................................................................153
4.6 References....................................................................................................155

Chapter 5. Protease-Modified Membranes for Studies of Protein
Structure: Limited Proteolysis and Rapid Digestion for HDX ..... 158
5.1 Introduction..................................................................................................158
5.2 Experimental ................................................................................................160
5.2.1 Materials, membranes and membrane holders .......................................160
5.2.2 Membrane modification with protease ..................................................160
5.2.3 Apomyoglobin HDX and membrane-digestion procedure ....................161
5.2.4 RHD3 expression and purification .........................................................161
5.2.5 Limited proteolysis of RHD 3 with either trypsin- or -chymotrypsinmodified membranes........................................................................................162
5.2.6 Mass spectrometry and data analysis .....................................................163
5.3 Results and discussion .................................................................................163
5.3.1 Preliminary study of HDX on apomyoglobin using digestion in a pepsinmodified membrane .........................................................................................163
5.3.2 Limited membrane proteolysis of RHD3 for structural information .....166
ix

5.4 Conclusions..................................................................................................175
5.5 Acknowledgement .......................................................................................175
5.6 References....................................................................................................177

Chapter 6. Summary and Future Work ........................................... 179
6.1 Research summary .......................................................................................179
6.2 Future work ..................................................................................................182
6.2.1 Study of the correlation of multiple phosphorylation events on p-tau and
-synuclein using membrane enrichment combined with limited digestion or
intact protein analysis ......................................................................................182
6.2.2 Phosphorylation mapping in tau isomers and multiple kinase events ...186
6.2.3 De novo sequencing of monoclonal antibodies using limited proteolysis
and a middle-down approach ...........................................................................187
6.2.4 Functional membranes for removal of affinity tags ...............................188
6.3 Summary of Future Work ............................................................................191
6.4 Acknowledgement .......................................................................................191
6.5 References....................................................................................................193

x

LIST OF TABLES
Table 2.1. Phosphopeptides identified from the mass spectrum (Figure 2.11) of the membraneenriched peptides from tryptic digests of -casein, -casein, and ovalbumin in loading buffer II.
The loading solution contained a 100-fold excess of BSA.. ......................................................... 83
Table 3.1. Phosphopeptides identified from membrane-enriched tryptic digests of the nuclear
extracts from TNF--treated THP-1 cells.. ................................................................................ 115
Table 3.2. Peptides identified using HPLC-ESI-MS/MS of 1 pmol of GST-IB digest.. ....... 120
Table 3.3. Peptides identified using HPLC-ESI-MS/MS of 1 pmol of GST-IB digest
phosphorylated by GRK5. .......................................................................................................... 121
Table 4.1. Apomyoglobin peptides identified from ESI-Orbitrap mass spectra of protein digested
in a pepsin-modified membrane.................................................................................................. 143
Table 4.2. BSA peptides identified from ESI-Orbitrap mass spectra of protein digested in a
pepsin-modified membrane.. ...................................................................................................... 149
Table 5.1. RHD3 peptides identified from MALDI and ESI-Orbitrap mass spectra obtained after
digestion in trypsin-modified and -chymotrypsin-modified membranes……………………..173

xi

LIST OF FIGURES
Figure 1.1. Schematic diagram of DESI ......................................................................................... 4
Figure 1.2. The cross section of the Orbitrap analyzer where the red line is ion trajectory. .......... 7
Figure 1.3. Peptide fragmentation notation based on the scheme created by Roepstorff and
Fohlman .......................................................................................................................................... 8
Figure 1.4. Simple representation of the bottom-up and top-down approaches to protein analysis
by MS.. ............................................................................................................................................ 9
Figure 1.5. Mechanism for kinase-catalyzed phosphorylation of serine. ..................................... 13
Figure 1.6. The left drawing represents the chelating bidentate binding mode between TiO2 and
salicylic acid, while the right drawing represents the bridging bidentate mode between TiO2 and
phosphate.. .................................................................................................................................... 21
Figure 1.7. Illustration of outcomes from ETD or CID (CAD) dissociation for short peptides,
peptides containing PTMs and large peptides .............................................................................. 26
Figure 1.8. The procedure to create a PLGA-PAH-PSS assembly on a nylon membrane. ......... 30
Figure 1.9. Three procedures to covalently link enzymes to an amine-containing surface via
different cross-linkers (a) glutaraldehyde, (b) DSC and (c) DSS. ................................................ 32
Figure 1.10. Conceptual drawing of HDX on a protein... ............................................................. 38
Figure 1.11. The workflow for obtaining protein structure information using HDX-MS ........... 40
Figure 2.1. Cartoon of a membrane containing TiO2 for phosphopeptide enrichment and a
miniaturized membrane holder with a syringe pump. .................................................................. 62
Figure 2.2. (a,b) TEM images showing the size distribution of TiO2 nanoparticles at two
different magnifications. (c) EDX analysis of the nanoparticles. ............................................... 69
Figure 2.3. Cross-sectional SEM images of nylon membranes before (a) and after (c) coating
with a PSS/TiO2 nanoparticle film. Images b and d show the EDX Ti maps for the images in (a)
and (c), respectively. ..................................................................................................................... 70
Figure 2.4. Calibration curve for the ICP-OES analysis of Ti. The square and triangle show the
absorbance of the diluted Ti loading solution before and after passing through the nylon
membrane………………………………………………………………………. ……71

xii

Figure 2.5. Breakthrough curve for the passage of 20 M phosphoangiotensin (in loading buffer
I, 0.1% TFA) through a 22 mm-diameter nylon membrane modified with a (PSS/TiO2)2 film. 73
Figure 2.6. Fraction of the phosphoangiotensin recovered from a loading solution after binding
and elution from a nylon membrane modified with a (PSS/TiO2)2 film.. .................................... 74
Figure 2.7. Calibration curve showing the ratio of H5 to D5 peptide signals from MALDI-MS
analysis. ......................................................................................................................................... 76
Figure 2.8. MALDI mass spectra of 1 L of a solution containing 100 fmol H5 peptide and 1.5
2

pmol digested BSA (a) before and (b) after passing through a small membrane (0.02 cm )
modified with (PSS/TiO2)2........................................................................................................... 77
Figure 2.9. MALDI mass spectra of (a) 500 fmol of D5 peptide internal standard with 333 fmol
of H5 peptide, and (b) 500 fmol of D5 internal standard plus 1 L of the eluate from enrichment
of 10 pmol H5 peptide into 30 L of elution buffer.. ................................................................... 78
Figure 2.10. (a) MALDI mass spectrum of 1 L of loading buffer I containing 150 fmol (each)
of digested -casein, -casein and ovalbumin. (b) Mass spectrum of the peptides collected from
100 L of the above solution by adsorption on a nylon membrane modified with (PSS/TiO2)2,
rinsing of the membrane with 100 L washing buffer I, and elution of the peptides with 10 L of
elution buffer. ................................................................................................................................ 80
Figure 2.11. (a) MALDI mass spectrum of 1 L of a loading solution containing 30 fmol (each)
of digested -casein, -casein and ovalbumin and 3 pmol of BSA in 260 mM urea. (b) Mass
spectrum of the phosphopeptides enriched from 500 L of the above loading solution by
adsorption on a (PSS/TiO2)2-modified nylon membrane, rinsing with 3 mL washing buffer II,
and recovery with 10 L of elution buffer. ................................................................................. 82
Figure 3.1. Cartoon of the NF-B pathway under the stimulation of TNF-After the receptor
outside the cellular membrane receives a signal (such as the cytokine TNF-. ......................... 92
Figure 3.2. Scheme of the protein interaction module-assisted phosphorylation.. ...................... 99
Figure 3.3. (a) MALDI mass spectrum of a digested FPLC fraction that contained approximately
800 fmol of GSK-3-p-tau complex along with associated proteins; (b) The corresponding mass
spectrum of peptides enriched from the above solution by adsorption on a TiO2-modified nylon
membrane and subsequent elution. ............................................................................................. 101
Figure 3.4. (a) MALDI mass spectrum of an FPLC fraction containing approximately 800 fmol
of digested tau protein and associated proteins. (b) The corresponding mass spectrum of
xiii

phosphopeptides enriched from the above solution by adsorption on a TiO2-modified nylon
membrane and elution. ................................................................................................................ 102
Figure 3.5. ESI-Orbitrap multistage CID-MS
386

n

of the [M+3H]

3+

precursor ion of
3

TDHGAEIVYKpSPVVpSGDTpSPR (m/z 819.34). (a) CID-MS/MS, (b) CID-MS of the -98
4

Da product ion from panel A (m/z 786.48), (c) CID-MS of the -98 Da product ion (m/z 753.98)
from panel B.. ............................................................................................................................. 104
Figure 3.6. ESI-Orbitrap HCD-MS/MS spectra of the [M+3H]

3+

386

precursor ion of

TDHGAEIVYKpSPVVpSGDTpSPR (m/z 819.34). .......................................................... 105

Figure 3.7. ESI-Orbitrap ETD-MS/MS spectrum of the [M+3H]

3+

386

precursor ion of

TDHGAEIVYKpSPVVpSGDTpSPR (m/z 819.34). ............................................................ 106

Figure
195

3.8.

ESI-Orbitrap

HCD-MS/MS

of

(a)

the

[M+2H]

2+
3+

pSGYpSSPGSPGTPGSR (m/z 778.00) and (b) the [M+3H]

407

precursor

ion

of

precursor ion of

HLpSNVSSTGSIDMVDpSPQLATLADESASLAK (m/z 1136.00). . ............................... 107

Figure 3.9. The WB analysis of p65 using primary polyclonal p65 antibody (C22B4). ............ 111
Figure 3.10. MALDI mass spectrum of a membrane-enriched digest of immunoprecipitated p65
and associated proteins.. ............................................................................................................. 112
Figure 3.11. MALDI CID-MS/MS spectra of (a) the [M+H]

+

+

precursor ion of

KEESEEpSDDDMGFGLFD (m/z 2029) and
(b) the [M+H]
precursor ion of
KEEpSEEpSDDDMGFGLFD (m/z 2109)................................................................................. 114
Figure 3.12. WB probe of phosphorylation of IB using phosphor- IB antibody............... 117
Figure 3.13 (a) TIC chromatogram of 1 pmol of a phosphorylated (by GRK5) IB digest (b)
The full-MS scan of peptides eluted at 18.71 min in (a), and (c) CID-MS/MS spectrum of the
3+
[M+3H] precursor ion at m/z 768.23 in (b)............................................................................. 118
Figure 4.1. Cartoon of controlled digestion using a membrane containing immobilized pepsin
and a miniaturized holder with a syringe pump. ......................................................................... 132
Figure 4.2. Bradford assay calibration curve for pepsin, and the absorbance of a pepsin solution
(0.4 mg/mL after a 5-fold dilution) before and after circulating through the membrane. .......... 138
Figure 4.3. Manually deconvoluted ESI-Orbitrap mass spectra of 0.1 mg/mL (6 M)
apomyoglobin digested with residence times of (a) 6, (b) 0.6, (c) 0.06 and (d) 0.02 s in a pepsinmodified membrane.. .................................................................................................................. 140
xiv

Figure 4.4. ESI-Orbitrap mass spectra of 0.1 mg/mL (6 M) apomyoglobin digested by a pepsinmodified membrane in (a) 6, (b) 0.6, (c) 0.06 and (d) 0.02 s...................................................... 141
Figure 4.5. ESI-Orbitrap mass spectra of (a) 0.1 mg/mL (6 M) apomyoglobin digested by a
pepsin-modified membrane for 6 s and (b, c) 100 g apomyoglobin digested by 5 g pepsin in
0.2 mL of 5% v/v FA. Digestion times are 5 min and 20 s for (b) and (c), respectively........... 142
Figure 4.6. Number (blue), average length (red), and average charge of peptides at the most
abundant peak (green) in ESI-Orbitrap mass spectra of 0.1 mg/mL apomyoglobin digested in a
membrane for 6 s, 0.6 s, 0.06 s and 0.02 s.. ................................................................................ 145
Figure 4.7. ESI-Orbitrap ETD-MS/MS spectrum of the [M+10H]

10+

precursor ion at 878.00..146

Figure 4.8. ESI-Orbitrap mass spectra of 0.1 mg/mL BSA (1.5 M in 0.06 M urea) digested in a
pepsin-modified membrane using (a) 2-s and (b) 0.05-s residence times .................................. 148
Figure 4.9. Peptides size distributions in ESI-Orbitrap mass spectra of 0.1 mg/mL BSA digested
in a pepsin-modified membrane for 2-s (blue, 32 peptides) and 0.05-s (red, 35 peptides). ....... 149
Figure 5.1. The workflow of HDX and membrane digestion. .................................................... 160
Figure 5.2. ESI-Orbitrap mass spectra of 6 M (a) deuterium-exchanged or (b) control
apomyoglobin digested by a pepsin-modified membrane (1-s residence time). (c) and (d) are
enlarged (x-axis) views of the selected peaks (red rectangles) from (a) and (b), respectively.. . 165
Figure 5.3. SDS-PAGE analysis of limited RHD3 digestions in a trypsin-modified membrane
(left) and an -chymotrypsin-modified membrane (right).. ....................................................... 168
Figure 5.4. MALDI mass spectra of 0.1 mg/mL (1.3 M) RHD 3 digested in residence times of
(a) 1 , (b) 0.2 and (c) 0.045 s by a trypsin-modified membrane, and (d) ESI mass spectrum of the
same sample as (c). ..................................................................................................................... 170
Figure 5.5. ESI-Orbitrap mass spectra of 0.1 mg/mL (1.3 M) RHD 3 digested in residence times
of (a) 1 , (b) 0.2 and (c) 0.045 s by a trypsin-modified membrane.. .......................................... 172
Figure 6.1. MALDI mass spectra of 2 pmol of p-tau digested for (a) 0.05 s in a trypsin-modified
membrane and (b) 16 h in solution. ............................................................................................ 185
Figure 6.2. Properties of -synuclein.. ....................................................................................... 185
Figure 6.3. Scheme of the protein interaction module-assisted phosphorylation... ................... 187

xv

LIST OF SCHEMES
Scheme 1.1. The workflow of the SIMAC strategy to enrich phosphorylated peptides. ............. 22
Scheme 1.2. (a) A procedure for phosphopeptides enrichment from cells prior to LC-MS/MS. (b)
The IMAC tip used for enrichment after the SCX in procedure (a). ............................................ 24
Scheme 1.3. The strategy for using PMO in the digestion of bacteriorhodopsin ......................... 36
Scheme 3.1 Workflow of the p65 phosphorylation study.......................................................... 110
Scheme 3.2. Workflow for the investigation of phosphorylation of IB catalyzed by IKKand
GRK5. ......................................................................................................................................... 116
Scheme 5.1. Strategy for limited RHD3 digestion in membranes containing trypsin and chymotrypsin............................................................................................................................... 167
Scheme 6.1. Comparison of enzymatic tag-removal in traditional and membrane-based work
flows............................................................................................................................................ 190

xvi

LIST OF ABBREVIATIONS
MS

Mass spectrometry

HDX

Hydrogen-deuterium exchange

MALDI

Matrix-assisted laser desorption/ionization

ESI

Electrospray ionization

DHB

2,5-Dihydroxy benzoic acid

HPLC

High-performance liquid chromatography

DESI

Desorption electrospray ionization

DART

Direct analysis in real time

m/z

Mass-to-charge

FT-ICR

Fourier transfer ion cyclotron resonance

RF

Radio frequency

TOF

Time of flight

LTQ

Linear trap quadrupoles

CID

Collision induced dissociation

CAD

Collision activated dissociation

ACS

American Chemical Society

PMF

Peptide mass fingerprinting

RP-HPLC

Reverse-phase high performance liquid chromatography

PTMs

Posttranslational modifications

ECD

Electron capture dissociation

ETD

Electron transfer dissociation

ATP

Adenosine-5’-triphosphate
xvii

Ser

Serine

Thr

Threonine

Tyr

Tyrosine

Arg

Arginine

SDS-PAGE

Sodium dodecyl sulfate polyacrylamide gel electrophoresis

WB

Western blotting

Lys

Lysine

His

Histidine

IMAC

Immobilized metal affinity chromatography

NTA

Nitrilotriacetate acid

TiO2

Titanium dioxide

MOAC

Metal oxide affinity chromatography

TFA

Trifluoroacetic acid

ASBMB

American Society for Biochemistry and Molecular Biology

SIMAC

Sequential elution from IMAC

SCX

Strong cation exchange

ACN

Acetonitrile

HCD

High-energy collision-induced dissociation

IMERs

Immobilized enzyme reactors

PLGA

Poly(L-glutamic acid)

PLL

Poly(L-lysine)

PAH

Poly(allylamine)

PSS

Poly(styrene sulfonate)

xviii

DSC

N,N’-disuccinimidyl carbonate

DSS

N,N’- disuccinimidyl suberate

PMO

Periodic mesoporous organosilica

Leu

Leucine

Phe

Phenylalanine

Ile

Isoleucine

Trp

Tryptophan

NMR

Nuclear magnetic resonance

UPLC

Ultra-performance liquid chromatography

bANc1p

Bovine mitochondrial ADP/ATP carrier isoform 1

CATR

Carboxyatractyloside

BA

Bongkrekic acid

BSA

Bovine serum albumin

ICP-OES

Inductively coupled plasma-optical emission spectroscopy

EDX

Energy-dispersive X-ray spectroscopy

GSK-3

Glycogen synthase kinase-3

TNF-

Tumor necrosis factor

THP-1

Human acute monocytic leukemia

IB

Inhibitor of kappa-B-alpha

IKK

Inhibitor of kappa-B kinase beta

GRK5

G-protein coupled receptor kinase-5

FPLC

Fast protein liquid chromatography

Met

Methionine
xix

cam

Carbamidomethyl

DTT

1,4-dithio-DL-threitol

TCEP

Tris(2-carboxyethyl) phosphine hydrochloride

NALS

Non-ionic acid labile surfactant

FA

Formic acid

RHD3

Root hair defective 3

SUMO

Small ubiquitin-like modifier

SEC

Size exclusion chromatography

His-tag

Polyhistidine-tag

xx

Chapter 1. Introduction
This dissertation describes the fabrication and use of functionalized porous membranes
for protein digestion and phosphopeptide isolation prior to mass spectrometry (MS) analysis.
More specifically, the research focuses mainly on TiO2-modifed membranes for phosphopeptide
enrichment and phosphoprotein characterization, and protease-modified membranes for limited
protein digestion. To provide context for the work, this chapter starts with an overview of MSbased proteomics and then reviews the importance of phosphoproteomics along with current
phosphopeptide enrichment and MS-based detection and characterization methods. With respect
to protease-modified membranes, a subset of enzymatic membrane reactors, this introduction
reviews the development of immobilized enzyme reactors in terms of immobilization methods,
available supports and applications. Additionally, this chapter describes two current MS-based
methods for studying protein structure, e.g., hydrogen-deuterium exchange (HDX) and limited
proteolysis, as these methods may benefit from the new protease-modified membranes. Finally,
the last section of this chapter outlines the dissertation.

1.1

Mass spectrometry for proteomics
With the completion of genome sequences for many plant and animal species, the focus

of much biochemical research has shifted to comprehensive studies of the proteins expressed by
these genes, i.e. proteomics. As proteins function like the building blocks of life, proteomics
research is vital to understand many life processes.[1] MS uses protein and peptide masses to
identify proteins and characterize their modifications, and because it is rapid, versatile and
potentially comprehensive, MS has become the most powerful tool in the proteomics area.[2]
Nevertheless, the complexity of the proteome makes comprehensive protein characterization a
1

daunting and as yet unfulfilled task. The desire to fully understand the proteome is continually
pushing the boundaries of instrumentation and methodologies, and new MS-based tools and
approaches appear regularly to extend the scope of proteomics applications.[3]
MS includes ionization, mass analysis, and complex data processing. This section briefly
describes basic concepts and new features in each of these areas. This knowledge is vital to the
later chapters in this dissertation, as MS-based analysis is the endpoint for the functional
membranes developed in this dissertation.

1.1.1 Ionization techniques
Since the invention of matrix-assisted laser desorption/ionization (MALDI)[4] and
electrospray ionization (ESI)[5] more than two decades ago, MS-based proteomics has improved
tremendously.[3, 6-7] Before MALDI and ESI, ionization of large peptides or proteins was
extremely challenging due to their thermal instability or low volatility. These two so-called “soft”
ionization techniques successfully ionize large species and turn them into gas phase ions without
significant degradation. The awarding of the 2002 Nobel Prize in Chemistry to the inventors of
ESI and MALDI in part reflects the impact of these innovations on MS-based proteomics.[8]

1.1.1.1

MALDI
The MALDI ionization process involves two steps. In the first step, the matrix, such as

2,5-dihydroxy benzoic acid (DHB) or -cyano-4-hydroxycinnamic acid, absorbs the laser energy
(typically at a wavelength of 337 or 355 nm). The ideal matrix will have good solubility in water,
strong optical absorption, a low molecular weight and facile vaporization, and additionally act as
a proton source. In the second step, the matrix transfers adequate energy to ionize the acidified
peptide or protein analytes and send them into the gas phase. The predominant ions are singly
2

+

charged, [M+H] . The MALDI mass spectrum is generated after one or a few laser shots, and
these spectra differ somewhat from shot to shot. In practice, a reliable spectrum with a sufficient
signal to noise ratio is the average of tens or hundreds of spectra.[9]

1.1.1.2

ESI
Unlike MALDI, the ESI ionization process generates ions from the solution containing

the analytes of interests. The liquid is dispersed into a fine aerosol and driven by the voltage (2-6
kV) between the capillary nozzle and mass spectrometer inlet. At the capillary tip, droplets
emerge from a Taylor cone in a well-defined physicochemical process.[10] In the subsequent
transition process, the solvent–analyte droplets gradually shrink due to the evaporation of solvent,
and when the charge repulsion force exceeds the surface tension the droplets split and reform
new ones in a process known as Coulomb fission. The analytes will have various charge states
when they reach the inlet of the mass analyzer.[11] Compared to MALDI, an obvious advantage
is that high charge states enable detection of larger molecular weight species under the same
analyzer m/z range. The continuous electrospray also makes on-line analysis achievable, so ESI
couples well to separation techniques like high-performance liquid chromatography (HPLC).
This coupling dramatically increases the throughput and sensitivity of detection. Furthermore,
improvement in Nano-HPLC or Micro-ESI are making the ESI ionization process even more
useful and practical.[12]

1.1.1.3

DESI and DART
Desorption electrospray ionization (DESI)[13] and direct analysis in real time

(DART)[14] are examples of emerging ionization methods. These techniques require little
sample preparation and can directly analyze gas, liquid and solid phase samples under ambient
3

conditions (Figure 1.1). The ionization takes place directly at the surface of samples ranging
from fruits, vegetables, blood, urine, and tissues.[13] The unique ambient properties of these
techniques may enlarge MS applications in proteomics, and I expect more application using
these methods in the future.

Figure 1.1. Schematic diagram of DESI. The solvent is influenced by high voltage and the sheath
gas direct sprays onto the sample surface. Desorbed ions are ejected into the ion transfer line,
with variation of anglesand to optimize collection. The figure is reprinted from reference
[13] with permission from the American Association for the Advancement of Science. For
interpretation of the references to color in this and all other figures, the reader is referred to the
electronic version of this dissertation.

4

1.1.2 Mass analyzers
A typical mass spectrometer contains a sample introduction and ionization source, a mass
analyzer, an ion detector and data processing electronics, but the mass analyzer is the core
component. The basic functions of the analyzer include storing or scanning ions and separating
them according to their mass-to-charge (m/z) ratios. There are four common strategies to
separate or store ions.[3] The first is frequency based analysis, e.g. Fourier transfer ion cyclotron
resonance (FT-ICR) and Orbitrap methods; the second is the stable trajectory of ions by specific
alternating and constant electric fields, and the representative analyzers contain quadrupoles; the
third combines the stabilizer principle and radio frequency (RF) and applies to ion traps; the last
principle is the travelling time of ions between the ion source and ion detector, and hence it is
termed as time of flight (TOF). The ion trap and Orbitrap deserve further details here, as they are
the two mass analyzers employed for all the results in this dissertation.

1.1.2.1

Linear ion trap
Thermo Scientific specifically terms the linear ion trap as the LTQ, or linear trap

quadrupoles. Compared to the traditional 3-D ion trap, one major advantage of the LTQ is that
the longer dimension of the ions’ traveling axis allows storage of more ions. The workflow and
mechanisms are almost the same for all ion traps. In detail, the ions created at the ion source
travel through a few sets of lenses and quadru/octopoles that help to concentrate and align the ion
beams. After ions enter the LTQ, they are stabilized through collision with a noble gas and
trapped at the center of the chamber by the trapping potential at both ends of the traveling axis.
Finally by increasing the amplitude of the RF field, the ions become unstable according to their
m/z value (low m/z first) and are ejected from the LTQ.[15] The LTQ has the advantages of easy

5

operation, low detection limits, rapid analysis, low maintenance, and variable resolution at a low
cost, so it is the most common mass analyzer.
The ion trap can also perform MS/MS analysis. The ions with specific m/z are first
selected from all ions trapped in the LTQ. Subsequently, the selected ions are excited to undergo
collisions with N2 or Ar gas. The collisions convert kinetic energy into internal energy that is
distributed to chemical bonds. Bond dissociation occurs when the distributed internal energy
exceeds the bond energy in a process known as collision induced dissociation (CID) or collision
activated dissociation (CAD).[16] The spectrum of fragment ions provides structural and
sequence information about the selected targets.

1.1.2.2

Orbitrap
Invented by Makarov in 2000 and commercially available from Thermo in 2005, the

LTQ-Orbitrap hybrid mass analyzer is gradually maturing as one of the leading analyzers for
proteomics research.[17-18] This hybrid mass analyzer retains the advantage of the LTQ and is
capable of high mass accuracy (1-10 ppm) and high mass resolution (up to 150,000).[18] The
ions trapped in the LTQ can transfer into the Orbitrap and are stabilized without magnetic or RF
fields. The ions are trapped in a quadru-logarithmic field, and both orbit radially around the axial
electrode (z) and oscillate along the axial electrode at different frequencies (), and the is
-1/2

proportional to (m/z)

(Figure 1.2). Similar to the FT-ICR, the analyzer measures the imaging

current induced by the oscillation and then transforms this current into a mass spectrum using a
fast FT.[17]

6

Figure 1.2. The cross section of
the Orbitrap analyzer where the
red line is ion trajectory. The
Figure is reprinted from reference
[17] with permission of the
American

Chemical

Society

(ACS).

Compared to the FT-ICR, the Orbitrap offers similar mass resolution and mass accuracy
at a lower price and maintenance cost, thus providing an alternative for analysis of intact proteins
or complex samples. However, the FT-ICR still has its unique properties of high mass range and
the capability to perform ion reactions in the ICR cell. In contrast, the Orbitrap works only as an
analyzer and needs the LTQ to perform the ion trapping and fragmentation or reaction.[2] In
conclusion, thanks to the Orbitrap, high mass accuracy and resolution facilitate the data analysis
process in peptide mass fingerprinting (PMF) and de novo peptide sequencing.

1.1.3 Proteomics approaches
With the availability of various advanced instruments, procedures to analyze proteins are
undergoing continuous refinements. However, all methods start with sample preparation aiming
to reduce complexity and to make the buffer compatible with the instrument. Chromatography
and two-dimensional gel electrophoresis are the most effective and widely used separation
methods prior to the MS analysis. In the instrument, MS data acquisitions have two major modes.
The first is the full-scan MS which detects and records a full spectrum at a given time, and the
7

n

other is a MS/MS (tandem MS) or MS scan, which analyzes the fragments (product ions) from
isolated ions (precursor ions). Known as data-dependent acquisition, the parameters (e.g. the
signal threshold, the number of selected ions and the selection criteria) used for a tandem MS or
n

MS scan usually depend on the results of full-scan MS.
Figure 1.3 shows the systematic labeling of the product ions that result from
fragmentation of a peptide. Cleavage of the amide bond will generate b ions (charge retained at
N-terminus side) and y ions (charge retained at C-terminus side). Similarly the a-x pair results
from cleavage of the C-C bond and the c-z pair stems from breaking of the N- Cbond.[19]
The product ions provide information on the peptide sequence. The analysis can occur with
proteolytic peptides, a bottom-up approach, or direct analysis of an intact protein, a top-down
approach (Figure 1.4).

Figure 1.3. Peptide fragmentation notation based on the scheme created by Roepstorff and
Fohlman.[19]
8

Fragment ions
of peptides

Proteolytic digests
(e.g. by trypsin)

Bottom-up
approach
Protein
Top-down
approach

Fragment ions
of protein

Figure 1.4. Simple representation of the bottom-up and top-down approaches to protein analysis
by MS. The bottom-up approach examines proteolytic digests and the fragmentation of the
digested peptides, whereas the top-down approach employs direct fragmentations of the intact
protein.

9

1.1.3.1

Bottom-up approach
To date, the bottom-up strategy is the most widely used method to analyze proteins in

complex mixtures.[20] This approach starts with digestion of the proteins. In the traditional
method, the proteins are first separated through two-dimensional gel electrophoresis, and the
proteolysis takes place in the gel prior to peptide extraction. The digests are then submitted for
MS analysis. Currently, most protein mixtures undergo in-solution digestion, the proteolytic
peptide mixture is separated through reverse-phase high performance liquid chromatography
(RP-HPLC), and the eluent is analyzed by ESI-MS on-line. This method greatly increases the
throughput of the process. For MS analysis, the full scan MS is first applied, and data-dependent
acquisition is triggered to obtain tandem mass spectra according to different settings. In the data
interpretation, both manual de novo peptide sequencing or database searching using algorithms
such as SEQUEST[21] can extract information from the tandem MS spectra. In database
searching, the actual tandem spectrum are compared with the predicted in silico fragmentation
results of peptides from the chosen set of proteins. Then the identified peptides will be used to
map the original proteins. Although this approach is effective, it has several drawbacks: the loss
of peptides during separation and ionization will decrease the probability of protein identification
and characterization; the peptide-protein mapping method will ambiguously assign peptides to
various isoforms; the proteolytic peptides cannot provide comprehensively identify
posttranslational modifications (PTMs). Ideally, analysis of the intact protein will solve these
potential problems, and I discuss that approach below.

10

1.1.3.2

Top-down approach
The top-down approach directly ionizes and fragments intact proteins instead of

proteolytic peptides.[20] Although the bottom-up approach remains the main tool for proteomics
analysis, the top-down approach is gathering more and more interest to obtain protein mass
information, differentiate isoforms, and characterize all PTMs at once.[22-25] Powerful mass
analyzers such as FT-ICR or Orbitrap instruments coupled with electron capture dissociation
(ECD)[26] or electron transfer dissociation (ETD)[27] make this approach feasible, as these
fragmentation methods efficiently break the peptide bonds and generate sufficient fragment ions
to provide sequence information. Nevertheless, for whole proteomics studies, the top-down
approach requires more material than the bottom-up approach and throughput of the top-down
method is still low.[28] Although the high mass accuracy and resolution in modern MS can
handle single protein or simple protein mixtures, the front-end chromatographic separation of
proteins is far behind RP-HPLC peptide separations, and this shortcoming limits the broad
replacement of the bottom-up approach for large-scale proteomics.[29]

1.1.3.3

Middle-down approach
In an effort to combine some of the assets of the above two approaches, middle-down

proteomics has emerged as a new way to examine multiple PTMs in a single long peptide (3 kD10 kD).[30-31] This method keeps the digestion from the bottom-up approach but attempts to
obtain large peptides through limited proteolysis. In particular, this approach was applied to
examine PTMs in Histones.[28, 32] In one histone H3 study, Kelleher and coworkers used the
endoproteinase Glu-C to release the tail peptide (residues 1-50) from the protein and performed
ECD fragmentation to obtain information on PTMs. The rationale is to efficiently fragment the

11

large peptide and get high sequence resolution from the fragments, as almost all methylation and
acetylation occur in the first 50 amino acid.[30]
So far this section has described some concepts and usage of MS in proteomics. Notably
the sample preparation is vital to achieve the desired results. My focus is to simplify the sample
preparation to facilitate the MS detection. Sample preparation is also vital in the development of
phosphoproteomics.

1.2

Phosphorylation: a key to protein function
PTMs significantly increase the proteome complexity and function in human cells, and

they also make the proteome a more challenging research area.[33-34] One category of the
PTMs is covalent modification with functional groups including phosphorylation, glycosylation,
methylation and acetylation. The second category of PTMs is the cleavage of one or more amino
acids from the original protein. Among the first category, protein phosphorylation is the most
common PTM and the most widely studied.[34] Around 500 protein kinases catalyze the transfer
of the phosphate group from adenosine-5’-triphosphate (ATP) to one or more substrate proteins ,
although ideally these protein kinases are chemo-specific.[35] The -position phosphate group is
the most common donor, and this covalent addition is reversible with the help of a phosphatase.
Reversible phosphorylation controls cell functions such as differentiation, apoptosis, and signal
transduction, consequently this phenomenon is involved in cell division, cell growth, and gene
expression.[33, 36-37]
In eukaryotic cells, phosphorylation occurs on serine (Ser), threonine (Thr) and tyrosine
(Tyr) residues, and early studies suggest that the ratio of modifications on these three sites is
1800:200:1.[38] In 2006, Mann and coworkers studied the phosphorylation events on Hela cells
stimulated with epidermal growth factor and revealed 6,600 phosphorylations on 2,244 proteins.
12

Their statistical results show that the relative percentages of phosphorylation on Ser, Thr and Tyr
are 86.4%, 11. 8% and 1.8%, respectively.[39] Thus, phosphorylation of serine is the dominant
modification. Figure 1.5 demonstrates the kinase-catalyzed mechanism in which serine takes a
-position phosphate from ATP to form phosphorylated serine. Thr and Tyr share the same
mechanism but with different kinases.
NH2
N
O

O

O

-B

N
OH

O

P

O

P

N

P

O

N

O
O

O

O

O

H
H

Mg2+

N
H

H

OH

H

H

O
O
O

Kinase

P

O

O

ADP

+
N
H
O

Figure 1.5. Mechanism for kinase-catalyzed phosphorylation of serine. The figure is adapted
from reference [43].

13

Given that the tertiary structure of proteins provides their unique properties,
conformational changes after phosphorylation are vital to mediate protein activity. Interestingly,
phosphorylation typically gives rise to changes from an inactive to an active protein mode, and
the opposite case is rare.[40] In the perspective of mechanisms, addition of phosphate will alter
not only the conformation in the region but also the local charge density, consequently both
factors will alter the protein microenvironment. As an example, the phosphorylated Ser14 of
glycogen

phosphorylase

charge-pairs

intramolecularly

with

arginine

(Arg69)

and

intermolecularly with Arg43. The interaction of two subunits drives an allosteric conformational
change to activate the enzymatic function of cleaving glucose-1-phosphate from glycogen.[40]
About one third of the proteins in humans are substrates for phosphorylation at some
stage in their life cycle,[41] but at any given time, the absolute amount of phosphorylated
proteins is much less than that of regular proteins. Moreover, for a certain protein, the amount of
the phosphorylated form may be lower than that of its unphosphorylated counterpart (low
stoichiometry), and different sites on a protein may be phosphorylated to different degrees. On
top of the above factors that complicate analysis of phosphorylation, because kinases and
phosphatases play their roles at the same time, phosphorylation is intrinsically dynamic and
transient.[42] The importance of phosphorylation and challenges in its detection are driving
continuous improvements in detection methods and instrumentation.[43]

14

1.3

Phosphoproteomics: detection of protein phosphorylation,
enrichment techniques and analysis strategy
As mentioned in the previous section, phosphorylation of protein side chains is one of the

most important PTMs and serves as a crucial mechanism for creating diversity in eukaryotic cells.
A fundamental understanding of regulatory pathways at the molecular level requires the
identification of the phosphorylation states in specific proteins. The following subsection
describes the historical methods for phosphorylation detection and the current leading MS-based
method. A subsequent subsection introduces several phosphopeptide enrichment strategies
employed prior to the MS analysis to facilitate detection, and the last subsection discusses MS
fragmentation methods.

1.3.1 Detection of protein phosphorylation
32

Initially, detection of the –[ P] phosphate transfer from ATP to the protein side chain
served as the primary method for examining phosphorylation.[44] In this technique, precipitation
of phosphorylated proteins using trichloroacetic acid separates them from a protein mixture, and
proves that the phosphorylations on Ser, Thr and Tyr are acid stable. After the protein washing, a
scintillation counter detects the

32

P if phosphorylation occurs. However, the localization of the

phosphorylation sites and determination of the stoichiometry are a challenge for this traditional
method. Moreover, this technique is more suitable for studying a targeted known protein than for
detection of global protein phosphorylation.
Based on sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE),
Western blotting (WB) is the most widely used analytical technique to detect proteins in a given

15

biological sample.[45] With appropriate phosphor-antibodies, WB is also the most common
approach to detect specific phosphorylation events.[46] Phosphotyrosine antibodies show the
most specificity for the phosphorylated residues, and binding of these antibodies does not depend
greatly on the surrounding amino acid sequence. Thus, antibody methods for detection of
phosphorylated tyrosine are highly developed.[47] However due to much less specific binding,
detection of phosphoserine and phosphotheronine with WB has developed more slowly. The
available phosphoserine and phosphothreonine antibodies rely on motif-based selectivity (e.g.
affinity to the RXRXXpT or pSXR sequence, where X is any amino acid).[46] In other words,
the antibodies will reveal phosphoserine or phosphotheronine only if they are present in select
sequences. Because of insufficient specificity, the antibodies for probing phosphoserine and
phosphotheronine are not widely used. Moreover, the success of this method depends on the
availability of the various phosphor-antibodies. This method is often a way to confirm a known
phosphorylation event rather than a discovery path to reveal new phosphorylation events.
Both 32P detection and WB are not high throughput techniques. From the perspective of
systems biology or global protein phosphorylation studies under a certain stimulus, the protein
complexity and abundance overwhelm the capabilities of these two detection methods. Currently,
MS is the leading technology for evaluation of the phosphoproteome [6, 38-39, 48-49] due to its
high sensitivity and throughput, and the capability to localize the phosphorylation site at specific
amino acids using tandem MS.[39]
The major goal of phosphoproteomics is to identify and quantify phosphorylated proteins,
and further to determine the location of the phosphorylated sites. This goal is normally realized
by examining the proteolytic peptides generated from a pure protein or a protein mixture.
Addition of 79.966 Da (phosphate group) to the known peptide mass is evidence for

16

phosphorylation. For an unknown sample, the signature indication is the loss of 97.995 Da
(trihydrogen phosphate) from Ser and Thr, or 79.966 Da (phosphate) from Tyr during a CID
process. The next step is to determine the peptide sequence and localize the phosphate groups at
n

certain amino acids. Localization relies on the product ions in the tandem MS or MS spectra,
where the product ions refer to the CID fragments of the selected peptides. The workflow is
straightforward, but there are many practical issues associated with the entire analysis such as
sample preparation (including phosphopeptide enrichment), peptide fragmentation and data
interpretation. Each step needs careful refinement to get accurate and reliable results for protein
phosphorylation.

1.3.2 Enrichment of phosphopeptides
Although reversible phosphorylation is ubiquitous in cells, the phosphoproteins are
generally present at relatively low abundance compared to the non-phosphorylated proteins
because of the intrinsically dynamic and transient nature of phosphorylation.[42] This low
abundance makes detection of phosphorylation extremely difficult in conventional MS analysis,
especially in complex mixtures. Additionally, two papers suggest that the non-phosphorylated
peptides have a higher ionization efficiency than their phosphorylated analogues.[50-51] Based
on the study of many synthetic phosphopeptides, Steen and coworkers disagree with the
statement that phosphopeptides have low ionization efficiencies.[52] However, this contradiction
may be due to the presence of lysine (Lys), Arg and histidine (His) in Steen’s analyzed
phosphopeptides.

These basic residues will increase the ionization efficiency when using

ESI.[53-54] Efforts have been made to enhance the signal intensity in MALDI-MS by adding

17

matrix additives like ammonium citrate[55] or phosphoric acid[56]. Nevertheless in complex
mixtures, phosphopeptide enrichment is a prerequisite for MS analysis.[57-58]
For decades, many strategies have been developed to carry out phosphopeptide
enrichment. The traditional biological process is immunoprecipitation, in which specific
antibodies capture phosphorylated species in a protein mixture. The most reliable antibodies are
only suitable for pTyr, so there are intensive studies of Tyr phosphorylation.[59-60] In contrast
the motif-based anti-pSer and anti-pThr antibodies limit studies of Ser and Thr phosphorylation.
[61-62]
Chemical derivatization is another important strategy for enriching phosphoproteins and
phosphopeptides. For example, -elimination of the phosphate groups present at Ser or Thr
residues followed by a Michael addition reaction can link a biotin-tag to the peptides or proteins
that originally contained the phosphate group.[63] Later, Zhou and co-workers introduced a
multi-step chemical modification using phosphoramidate chemistry for all phosphorylated
residues.[64] Unfortunately, the general drawback of chemical derivatization is that potential
side reactions and conversion yields below 100 % result in both sample loss and increased
complexity.[65]
The most common phosphopeptide enrichment strategy is affinity chromatography. First
introduced in 1975, immobilized metal affinity chromatography (IMAC) initially aimed to
fractionate proteins based on the coordination interaction between a metal ion and the imidazole
group on histidine.[66] Metal chelators such as iminodiacetic acid and nitrilotriacetate acid
(NTA) link the affinity metal ions to the chromatographic support.[67] Today, IMAC columns
are one of the primary tools for purifying tagged recombinant proteins. Introduced by Andersson
3+

and Porath in 1986,[68] IMAC using Fe

complexes (IMAC-Fe) has been the most common

18

method to enrich phosphopeptides. IMAC-Fe relies on hard acid-hard base interactions between
positively charged metal-ligand complexes and specific negative functional group such as
3+

3+

3+

phosphates. Other candidate metal ions include Al ,Ga ,Co

4+

and Ti ,[69] and this technique

is widely used in silica column[70], magnetic beads[71] and modified MALDI plates.[72]
In the past, titanium dioxide (TiO2) served as a stationary phase to adsorb organic
phosphate or phospholipids.[73-74] More recently, TiO2 has been widely used as an alternative
to IMAC for phosphopeptide enrichment.[73-76] At acidic pH, TiO2 has a positively charged
surface that selectively adsorbs phosphorylated species,[77] and the adsorbed phosphopeptides
can be easily eluted at basic conditions.[78] This technique belongs to the large category of metal
oxide affinity chromatography (MOAC).[79-80] Unlike IMAC, which requires frequent
recharging (loading of the metal ion) before use, the MOAC material themselves (TiO2 or ZrO2
beads) can be the packing material for a column.[2] Moreover TiO2 has been reported to be more
effective than IMAC in the specific enrichment of phosphopeptides.[80-88] Furthermore TiO2
columns already have a number of online applications in phosphopeptide enrichment, and this
online approach greatly improves throughput and enhances global phosphorylation studies.[80,
85, 89]
Unfortunately, both IMAC and MOAC suffer from unwanted binding of highly acidic
peptides. Thus peptides containing multiple glutamic acid and aspartic acid residues frequently
co-purify with phosphopeptides.[79] O-methyl esterification can block the acidic residues and
enhance the specific phosphopeptide binding.[34] However, incomplete esterification
complicates MS analysis.[90] More recent studies suggest that excluders such as DHB[79, 81,
19

91-92] can block the non-specific binding sites without influencing the phosphopeptide
binding.[93-94] Jenson and Larsen suggest that compounds with different coordination
geometries have affinity towards different sites on TiO2. Figure 1.6 shows the coordination
modes between TiO2 or salicylic acid and phosphate. The bridging-bidentate binding between
phosphate and TiO2 does not compete with sites for the chelating-bidentate binding between
substituted carboxylic acid and TiO2. However, the chelate-bidentate binding does affect
adsorption of acidic peptides. A list of excluders in order of inhibiting efficiency includes: DHB
~ salicylic acid ~ phthalic acid > benzoic acid ~ cyclohexane carboxylic acid > phosphoric acid >
trifluoroacetic acid (TFA) >acetic acid.[79] Careful selection of loading, washing and elution
buffer will improve the selectivity and efficiency of phosphopeptide binding.[81, 89]
Another empirical rule is that TiO2 prefers mono-phosphopeptides over multiphosphopeptides, whereas IMAC selectively enriches multi-phosphopeptides.[95-96] I presume
that this phenomenon reflects incomplete elution of multi-phosphopeptides from TiO2 rather than
a lack of binding of these species. In addition IMAC sometimes suffers from insufficient
binding capacity, and the multi-phosphopeptides likely occupy binding sites rather than monophosphopeptides. Multi-phosphopeptides readily elute from IMAC resins, and TiO2 usually has
no binding capacity problem. Liang et al. showed a binging capacity of 300 mol of phenyl
phosphate per gram of cross-linked TiO2 nanoparticles.[97]

20

Figure 1.6. The left drawing represents the chelating bidentate binding mode between TiO2 and
salicylic acid, while the right drawing represents the bridging bidentate mode between TiO2 and
phosphate. The figure is reprinted from reference [79] with permission from the American
Society for Biochemistry and Molecular Biology (ASBMB).
Due to their intrinsic selectivities for mono-phosphorylated and multi-phosphorylated
peptides, respectively, TiO2 and IMAC are sometimes used together to achieve complementary
information.[95-96] Thingholm and coworkers developed a sequential elution from IMAC
(SIMAC) strategy to increase the comprehensiveness of phosphopeptide enrichment. Scheme 1.1
illustrates their SIMAC workflow to enrich phosphorylated peptides. They first employ IMAC
beads in GELoader tips to enrich phosphorylated peptides. After incubation for 30 min and
washing, acid elution (1 % TFA, pH 1.0) is applied to elute the mono-phosphopeptides. Peptides
that passed through the column in the loading, wash and acid elution are enriched by adsorption
on TiO2. The eluent from TiO2 (primarily mono-phosphopeptides) and a basic eluent (NH4OH,
pH 11.3) that removed phosphopeptides remaining on the IMAC resin (primarily multiphosphopeptides) are analyzed by MALDI MS or LC-ESI MS/MS. In a phosphoproteome study
21

of 120 g of protein from human mesenchymal stem cells, compared to an optimized TiO2
enrichment method, the SIMAC identified two-fold more total phosphopeptides, including a
three-fold increase in multi-phosphopeptides.

Mixing IMAC beads
and peptide samples
IMAC beads
Non-phosphorylated peptide
Phosphorylated peptide

IMAC flow-through
and wash

TiO2 chromatography

TiO2 elutent

Parking beads onto
GLEloader tips

Acid elution
1% TFA pH 1.0

Basic elution
NH4OH pH 11.3

TiO2 chromatographyb

TiO2 elutent

MALDI MSa LC-ESI MS/MSb
a

Scheme 1.1. The workflow of the SIMAC strategy to enrich phosphorylated peptides. Step is
b

performed only in a simple peptide mixture and Steps are performed only in the complex
peptide mixture. The figure is reprinted from reference [95] with permission from the ASBMB.
22

For even larger scale phosphorylation studies (e.g. over five thousand phosphorylation
events), the IMAC and/or MOAC enrichment are not adequate to reduce the sample complexity
while maintaining all valuable information. Similar to RP-HPLC, strong cation exchange (SCX)
chromatography fractionates the complex sample and alleviates the workload for later
enrichment steps. At pH 2.7, the phosphate group will bring a negative charge to modified
peptides, so compared to non-phosphorylated peptides, phosphopeptides should elute in early
fractions of the SCX eluate. The SCX approach divides one extremely complex sample into tens
of moderately complex samples, on which either IMAC or MOAC have reasonable enrichment
performance. Scheme 1.2 demonstrates the procedure for phosphopeptide enrichment and
analysis from a yeast lysate. In the protocol, the protein extracts from yeast cells are first reduced,
alkylated and digested with trypsin in solution. The SCX fractionates the peptides mixture into
12 tubes. Each fraction undergoes desalting and enrichment by IMAC and eventually analysis by
LC-MS/MS.[98] With this SCX/IMAC enrichment protocol to explore the phosphoproteome in
several systems, Gygi and Villé found 5,500 and 13,000 phosphorylation events in mouse and
n
Drosophila embryo characterizations, respectively.[99-100]

23

S.cerevisiae

% Phosphopeptides

Cell lysis
IMAC/desalting
Proteins
SCX

% Phosphopeptides

Reduction
Alkylation
Trypsin-digestion

Add solution

IMAC resin
C18 discks

LC-MS/MS

Scheme 1.2. (a) A procedure for phosphopeptides enrichment from cells prior to LC-MS/MS.
Extracts for yeast are reduced, alkylated and digested with trypsin in solution. The SCX
fractionates the peptide mixture into 12 tubes. Each fraction undergoes desalting and enrichment
by IMAC and eventually analysis by LC-MS/MS. (b) The IMAC tip used for enrichment after
the SCX in procedure (a). The phosphopeptides trapped on the IMAC resin will be first eluted
onto the C18 disks without acetonitrile (ACN), next rinsing will desalt the sample, and finally
phosphopeptides are eluted by organic solvent. The figure is reprinted from reference [98] with
permission from the Nature Publishing Group.

1.3.3 MS Analysis Strategy
Both fragmentation methods and gas phase ion chemistry play a vital role in the
characterization of phosphopeptides.[101] In the conventional bottom-up approach, the
phosphopeptides are traditionally fragmented by CID. The tandem MS identifies certain species
as phosphopeptides based on the neutral loss of 98 Da or 80 Da. However these particularly
neutral loss fragments dominate the tandem mass spectrum so little sequence information is
24

available. A data-dependent acquisition mode supplements this incomplete information, as MS3
3

is triggered at the dominant peak in tandem MS. The MS spectrum contains the sequence
information for the peptide.[2, 101]
Although this is a universal approach, some gas phase ion chemistry will influence the
results under certain circumstance. When using MALDI as the ion source and an ion trap as the
analyzer, Palumbo et al. observed phosphate rearrangement and competing loss phenomena in
some cases. From studies on synthetic phosphopeptides, they conclude that phosphate
rearrangement happens at a high probability when the number of basic residuals (Lys, Arg and
His) is larger than the peptide charge (usually one in MALDI ionization). There is also a
potential for competing losses of 98 Da from either the precursor ion or from the b- and y-type
product ions. (either directly as 98 Da (H3PO4) from a phosphorylated Ser or Thr residue, or as
the combined losses of 80 Da (HPO3) from a phosphorylated Serine or Threonine residue and 18
Da (H2O) from a non-phosphorylated Ser or Thr residue). Both events result in ambiguity in
assigning the sites of phosphorylation within the peptide sequence.[102]
As an alternative fragmentation method to characterize the phosphorylation sites and
peptide sequence, high-energy collision-induced dissociation (HCD) is carried out exclusively in
another cell provided with the LTQ-Orbitrap instrument.[103] Based on the same collision
mechanism as CID, the HCD uses increased radio frequency voltage (2500 V compared to 1500
V) for containing more ions in the trap. This modification leads to more fragmentation in the cell,
and production of more b- and y-type product ions in addition to immonium ions. The more
comprehensive fragmentations facilitate the de novo sequencing and localization of
phosphorylation.
25

Different from the collision of positively charged peptides with a noble gas, gas-phase
ion/ion chemistry is another path to induce fragmentation.[26-27] In the ETD process, an anion
with low electron affinity, such as fluoranthene ion, donates an electron to the multi-charged
peptide cation to generate one hydrogen radical that initiates peptides fragmentation. The largest
difference is the fragmentation pathway, where the CID fragments first at the lowest energy bond
and then the amide bonds, ETD favors cleavage of the C-N bond and the labile modification
will remain on its residue.[104] As a result, the localization of phosphorylation is straightforward
and clear. Figure 1.7 is an analogy of the outcome of the dissociation from CID (CAD) or ETD
for short peptides, peptides with PTMs and large peptides.

Figure 1.7. Illustration of outcomes from ETD or CID (CAD) dissociation for short peptides,
peptides containing PTMs and large peptides. ETD is effective for the three types of peptides,
26

while CID works well for short peptides but less effectively for peptides containing PTMs and
large peptides. The Figure is reprinted from reference [104] with permission from the ACS.
One of the inventors of the ETD technique, Coon, and his coworkers recently conducted
a phosphoproteome study on the human embryonic stem cell and revealed 8087 unique
phosphopeptides by ETD and 3868 unique phosphopeptides by CID.[105] Although the ETD
approach is superior to the CID method in terms of peptide identification efficiency and
phosphorylation assignment, many factors such as precursor ion charge, precursor ion
mass/charge ratio and identity of peptides will great influence the results in both methods.[106]
Phosphoproteome studies should employ both fragmentation methods if possible.
Inspired by these pioneering studies, this dissertation (chapters 2 and 3) introduces a
membrane-based TiO2 enrichment technique and its investigation of the phosphorylation of tau
protein.

1.4

Immobilized enzyme reactors
Immobilization of enzymes on a support already has a history of almost 100 years.[107]

In recent developments, immobilized enzyme reactors (IMERs) have found applications in the
areas of digestion, catalysis of small-molecule reactions (e.g. glucose oxidase) and enzyme
inhibitor screening.[108-109] This section will focus on the contribution of IMERs to proteomics,
specifically the usage of immobilized proteases such as trypsin and pepsin to digest proteins
prior to their MS analysis, as this is related to the work in chapters 4 and 5.
As emphasized above, sample preparation is a vital step in MS-based proteomics. For a
bottom-up approach, an efficient and effective proteolysis will yield high quality peptidecontaining samples for successful protein identification and characterization.[108, 110] IMERs
27

containing proteases usually contain a high enzyme density to catalyze digestion in a few
seconds, and this fast digestion greatly increases the throughput for proteomics studies compared
to the traditional 16-h in-solution digestion. Other assets of digestion in IMERs include minimal
autolysis of the protease, reusability and on-line automation.[108, 110] The immobilization
process stabilizes the proteases and enhances their tolerance to high temperatures, pH variations
and organic buffer.[109] Most column-based IMERs can directly couple to HPLC-MS
instruments to reduce sample handling, increase sensitivity and decrease analysis time.[111] The
choice of immobilization methods, supports and enzymes plays an important role in tailoring
IMERs for a particular purpose, and the following subsections discuss these three variables.
Unfortunately, there is no universal protocol for creating effective IMERs.

1.4.1 Immobilization methods
The major techniques to immobilize enzymes are either chemical linkage with covalent
bonds or physical adsorption that relies on hydrogen bonds and hydrophobic or ionic interactions.
The main goal is to trap the enzymes on the support while retaining their activity. Among the
available anchoring methods, hydrophobic adsorption to the support is the simplest but also the
least robust. Lee et al. immobilize trypsin onto a polyvinylidene fluoride membrane via
hydrophobic interactions.[112] They successfully incorporate the miniaturized trypsin-modified
membrane in a common capillary fitting and perform digestion prior to online nano-ESI-MS.
The method requires very little sample consumption (<5 fmol) and achieves low detection limits
-8

(10 M). However replacement of the protease by hydrophobic proteins or leaching of protease
in the presence of surfactants used to increase the protein solubility will limit the lifetime of the
IMER.[111]

28

Proteins adsorption via electrostatic interaction is another versatile method suitable for
enzyme immobilization on charged substrates.[113-117] Moreover the bound enzymes
demonstrate similar or higher activity compared to the free enzyme in solution.[113-115]
Bhattacharyya and coworkers use layer-by-layer-assembled films to capture enzymes in
membranes. They first covalent attach either poly(L-glutamic acid) (PLGA) or poly(L-lysine)
(PLL) to the surface and pores of the membrane substrate.[113] Then they electrostatically
adsorb poly(allylamine) hydrochloride (PAH) and poly(styrene sulfonate) (PSS) to create PLLPSS-PAH or PLGA-PAH-PSS assemblies, which have positive and negative surface charges,
respectively. Figure 1.8 shows the three steps to create a negatively charged PLGA-PAH-PSS
assembly. As expected, the adsorption is much greater when the protein charge is opposite to the
charge on the substrate than when the two charges have the same sign. Although this electrostatic
immobilization method is facile and versatile, leaching or replacement of the enzyme is possible
when the buffer has a high ionic strength or incoming proteins are highly charged.

29

Figure 1.8. The procedure to create a PLGA-PAH-PSS assembly on a nylon membrane. The
PLGA is first covalently attached to the nylon membrane, and positively charged PAH absorbs
to the negatively charged PLGA through electrostatic interactions. Similarly, the PSS attaches to
the PAH and leaves the surface negatively charged. This Figure is redrawn according to
reference [113] with permission of the ACS.
Compared to adsorption, covalent binding of enzymes to a substrate will provide more
robust attachment. The most common covalent binding method includes activation of supports
containing amino group with glutaraldehyde, a homo-functional cross-linker (Figure 1.9a).[109]
The glutaraldehyde reacts with the amine on the support via one aldehyde and leaves the other
aldehyde ready to connect to the protein N-terminus or a free amine of a lysine residue. To
prevent potential hydrolysis, the labile Schiff base is further reduced by sodium borohydride in
some cases.[118] Other strategies employ N,N’-disuccinimidyl carbonate (DSC, Figure 1.9b) or
30

N,N’- disuccinimidyl suberate (DSS) as a linker (Figure 1.9c). With DSS a six-carbon chain
spaces the enzyme from the surface, and the enzyme linking occurs via an amide bond.[119] The
covalent method provides stable immobilization, but the enzyme concentration in the
immobilization solution must be high, and the reaction may require many hours.[118, 120-121]
Moreover, a recent review also discusses the concern of enzyme-activity loss during the covalent
immobilization reaction.[122] A number of other reactions are also available for covalent
immobilization. [109]
In summary, all enzyme-immobilization methods developed so far are effective but have
some drawbacks. The choice of a specific method may need to balance the project goals with
aspects that can be sacrificed, such as short reaction time, low starting material concentrations or
partial loss of enzyme activity.

31

Figure 1.9. Three procedures to covalently link enzymes to an amine-containing surface via
different cross-linkers (a) glutaraldehyde, (b) DSC and (c) DSS. The figure is reprinted from
reference [109] with permission from the Elsevier .

1.4.2 Immobilization supports
The basic criteria to choose the support are high surface area, good mechanical strength
and stability over the necessary range of pH values and temperature. For most applications both
polymeric and inorganic materials are acceptable. Supports are already available in a wide range
of geometries including beads, monoliths, nano/micro particles, microfluidic channels and
membranes. Below I describe a few examples of specific supports for enzyme immobilization.
32

Magnetic microspheres are attractive supports because they can be readily recovered
from complex mixtures using a magnetic field. Zhang and coworkers employ a one-step reaction
to bind trypsin to silica-coated magnetic microspheres via a 3-glycidoxypropyltrimethoxysilane
linker.[123-124] In addition to their magnetic properties, these microspheres adsorb microwave
radiation to promote digestion in solutions containing the trypsin-modified beads. A 15-s
microwave-enhanced digestion was equivalent to or better than the traditional 16-h digestion
with free trypsin.[123]
Monoliths have emerged as attractive enzyme supports in recent years. Both silica and
organic polymers can form monolithic disks, columns and capillaries.[111] In addition to
chemical and physical stability, the high surface area and low backpressure of such porous
monoliths make them attractive for diverse enzyme-based applications, including coupling to
HPLC-MS for online reactions.[120] Svec and coworkers successfully immobilized trypsin and
endoproteinase LysC onto a poly-(glycidyl methacrylate-co-ethylene dimethacrylate) monolith.
They demonstrate that hydrophilization via photografting of poly(ethylene glycol) methacrylate
greatly reduces nonspecific adsorption of hydrophobic proteins and peptides, and later they apply
this enzyme reactor to the digestion of immunoglobulin G. A 6-min digestion is similar to insolution digestion for 24 h.[120]
Microfluidic devices are appealing for online biological and chemical analyses with
minimal sample consumption.[125] Liu et al. incorporate proteases and charged polysaccharides
into a microchip using layer-by-layer assembly. They achieve a detection limit as low as 0.1
ng/L with cytochrome c digests, and the microchip rector digests femtomoles of protein per
analysis within seconds.[115] They conclude that the high enzyme density in the micro-channel
increases digestion rates three orders of magnitude relative to in-solution digestion.

33

Commercial polymer membranes serve as filters in many applications, but they are also
excellent supports for enzymes.[126] Their large surface area to volume ratio leads to a high
density of immobilized enzymes, and short radial diffusion distance in pores will allow enzymesubstrate reactions without diffusion limitations. Furthermore, the small membrane thickness
(~100 m) enables convective transport with a simple syringe pump. Our previous group
members, Xu and Wang developed membrane-based digestion in which a 6-s digestion of BSA
leads to 84 % sequence coverage in MALDI-MS. This sequence coverage is significantly higher
than the 71 % obtained after in-solution digestion for 16 h.[114] Moreover trypsin-modified
membranes still function in the presence of 0.05 wt% SDS, whereas trypsin in solution does not
catalyze digestion under similar conditions. This is another evidence that immobilization
enhances enzyme stability. In summary enzyme immobilization on a variety of supports is
creating exciting opportunities for IMERs in proteomics.

1.4.3 Immobilized trypsin and pepsin, reactors for protein digestion
Although there are a number of proteases applicable to digestion for proteomics, trypsin
and pepsin are the most readily available. In the bottom-up approach, trypsin is the protease of
choice due to its low cost, predictable selectivity, and efficient performance.[127] Trypsin
catalyzes protein cleavage at the C-terminus of Lys and Arg residues, unless the following amino
acid is proline. Usually, new digestion methods pursue rapid and complete digestion, and the
studies described in the previous subsection all achieved this goal. However, the digestion
mechanism is also important for controlling the process.
Liu and coworkers use a nanoporous material to confine the protease-substrate reaction
and find that confinement increases the digestion efficiency. BSA and myoglobin digestion with
such trapping materials yield more peptides than traditional in-solution digestion. Using MS
34

fingerprinting, they identify 293 proteins after nanopore digestion of a normal human liver
cytoplasm sample, whereas analysis of an in-solution digest leads to identification of only 100
proteins.[128] In a subsequent study, they propose that in a confined environment partially
digested peptides compete with the intact protein for digestion sites. Cleavage of the peptides is
more likely than proteolysis of the intact protein, so complete digestion occurs more readily than
digestion of a protein into a few pieces.[129] Very recently, Liu’s group invented a periodic
mesoporous organosilica (PMO) material for membrane protein digestion.[130] Due to the
hydrophobicity and structure rigidity, membrane proteins resist proteolysis. Scheme 1.3
describes the strategy for using the PMO to digest bacteriorhodopsin. This amphiphilic PMO can
adsorb and trap hydrophobic membrane proteins into its pores and later concentrate trypsin from
aqueous solution into pores and promote proteolysis. To prove the general utility, they
successfully apply this material for identification of membrane protein extracted from mouse
liver. [130]

35

Scheme 1.3. The strategy for using PMO in the digestion of bacteriorhodopsin. The PMO first
extracts the membrane protein from methanol into its pores, and after centrifugation the PMO is
immersed into a trypsin-containing solution to adsorb trypsin for confined digestion. Finally, the
peptides are eluted for MS analysis. The figure is reprinted from reference [130] with permission
from the ACS.
Pepsin is also a relatively inexpensive protease, but it is less specific than trypsin.
Because pepsin shows a preference for cleavage after most hydrophobic amino acid, many amide
bonds are potential cleavage sites.[131] A statistical analysis from 39 peptic protein digestions
demonstrates that the probability of N-terminus cleavage after leucine (Leu) and phenylalanine
(Phe) is greater than 40 %, whereas other residues such as isoleucine (Ile), Tyr and tryptophan
(Trp) are less vulnerable.[132] Unlike trypsin, pepsin shows optimal activity under acidic
conditions (pH 1-2) that may partially denature proteins and, in particular, help to solubilize
36

membrane proteins. Zhang et al. couple a pepsin-modified column to a SCX-RPLC-ESI-MS/MS
system to improve the digestion and identification of membrane proteins. In their study they
annotate 91 out 235 identified proteins as membrane proteins with transmembrane domains.[133]
Wilson and coworkers also immobilize pepsin in a microfluidic reactor to perform digestion and
on-chip electrospray ionization. They achieve high sequence coverage with peptic digests of
myoglobin, ubiquitin and bovine serum albumin using a digestion time of 4 s.[134] The Forest
group finds that apomyoglobin digestion with a pepsin/rhizopuspepsin-containing column gives
more complete proteolysis than conventional solution-phase digestion.[135] These studies are
particularly relevant to the use of pepsin for digestion in HDX studies, which I describe in the
next section.

1.5

Hydrogen-deuterium exchange and limited proteolysis for MS
As mentioned in the protein phosphorylation discussion, changes in protein structure or

conformation and protein-protein interactions are key factors controlling protein function. X-ray
crystallography and high-resolution nuclear magnetic resonance (NMR) remain the dominant
tools for studying protein structure and dynamics.[136] However, these techniques are timeconsuming and require either protein crystals, which are often not available, or highly pure
protein.[137]
In the 1950s, Kaj Ulrik Linderstrø
m-Lang first explored the deuterium exchange
phenomenon in proteins.[138] Remarkably, under physiological conditions the hydrogen on the
amide bond (except Proline) rapidly exchanges with deuterium when an unfolded region is
exposed to D2O, but in a folded region or areas with strong hydrogen-bonding, the hydrogen on
the amide bond exchanges at a much slower rate or even not at all (Figure 1.10).[139] By

37

investigating which areas of a protein undergo deuterium exchange, one can generate lowresolution structural information. MS is the natural tool to analyze the mass increases due to
HDX, and the first HDX-MS studies on proteins appeared in 1991.[140] HDX-MS research is
now an important tool for exploring regional protein structure and dynamics, with expanding
applications.[137, 139, 141] The following subsections describe HDX-MS applications using
top-down and bottom-up approaches for analysis.

Figure 1.10. Conceptual drawing of
HDX on a protein. The green balls
represent the amide hydrogens, and
the

red

balls

stand

exchanged deuterium

for

the

after the

protein is exposed to D2O.

The

hydrogens on the loops and some
unfolded helices exchange with
deuterium rapidly. The hydrogen in
the folded or protected areas has
very slow or negligible exchange
rates. The figure is reprinted from
reference [136] with permission of
the ACS.

38

1.5.1 HDX-MS using the top-down approach
After allowing HDX for a given time, the protein can directly undergo electrospray
ionization and fragmentation by ECD. Konermann and coworkers demonstrate the success of
this approach on a small protein, horse myoglobin.[142] They compare the deuterium exchange
state (where and how many deuterium are in the amide bonds) between the native
holomyoglobin and the apomyoglobin without the prosthetic heme group. ECD fragmentation
gives a resolution of less than two amino acids and result shows that the F-helix is unfolded in
the apomyoglobin and the G- and H-helix are also partially denatured. The conclusion is based
on greater deuterium exchange in those region in the apo-form compared to the native form, and
the result is consistent with previous NMR data.[143] The ECD fragmentation greatly reduces
the H/D scrambling that may occur during the CID process. In CID, intermolecular proton
migration results in ambiguous assignments of H/D on the amide bonds.[144] Another common
concern is back-exchange, where the exchanged deuterium will return to hydrogen if the solvent
changes back to H2O, but that issue does not exist in this study due to the on-line procedure that
requires no buffer exchange. However, this method only applies to proteins with a limited size.
Larger proteins do not undergo efficient fragmentation, so the amino acid resolution will be very
low.

1.5.2 HDX-MS using the bottom-up approach
Most HDX studies employ a bottom-up approach, in which the HDX is first quenched by
lowering the pH to 2.5 and the temperature to 0 º In this environment, the exchange rate is five
C.
orders of magnitude slower than that at physiological conditions.[137] John Engen employs an
effective protocol for analysis of HDX: After HDX and quenching, the protein undergoes rapid
39

peptic digestion followed by ultra-performance liquid chromatography (UPLC) and ESI-MS or
ESI-MS/MS by ETD (Figure 1.11). The main goal is to shorten the time between the quenching
and MS analysis to reduce the chance of back exchange and reflect only the exchange in the
native state. High amino acid resolution in ESI-MS relies on the overlap of peptides generated
from peptic digestion.[137, 145] Pepsin is the enzyme of choice for the low pH condition, and
immobilized-pepsin reactors enhance the digestion efficiency and shorten the digestion time to
enhance amino acid resolution and prevent back-exchange. Thus, further improvement in online immobilized-pepsin reactors is important to facilitate HDX studies.[135, 146]

Figure 1.11. The workflow for obtaining protein structure information using HDX-MS. In the
old pathway, the protein is labeled in D2O and quenched by acid manually, the digested peptides
are separate by HPLC on ice and analyzed by ESI-MS, and data is interpreted by hand. In the
new protocol, the robotic automation precisely controls the labeling and quenching, and UPLC
with refrigeration reduces the back exchange during peptide separation. Multiple MS analysis
tools are available for different purposes, and the data processing is achieved with automated
algorithms. The figure is reprinted from reference [136] with permission of the ACS.

40

1.5.3 HDX-MS using multiple acidic proteases
In the bottom-up approach, the spatial resolution for a high quality deuterium-exchange
determination depends on a nearly complete, but overlapped digestion. Forest and coworkers
employ both pepsin and rhizopuspepsin to improve the digestion[135] and study the bovine
mitochondrial ADP/ATP carrier isoform 1 (bANc1p), which plays an important role of
catalyzing the transport of ADP/ATP across the mitochondrial inner membrane in energy
metabolism. In a conformational dynamics study of this protein, they discover that two toxin
blockers (carboxyatractyloside or CATR, and bongkrekic acid or BA) induce different
conformational changes when they form CATR-bANc1p and BA-bANc1p. The CATR-bANc1p
complex result from HDX-MS agrees with their three-dimensional structure of CATR- bANc1p,
and this agreement validates their method. The structure of BA-bANc1p is unknown, so the
findings about BA- bANc1p from HDX-MS provide important information for the structure and
mechanism.[147]

1.5.4 Limited proteolysis for protein structure elucidation
Limited proteolysis, incomplete digestion of a non-denatured protein, is another
traditional method to probe protein structure, but it is not often combined with MS. This
technique is particularly useful for identifying disordered regions of a protein, removing the
loose and flexible regions from a protein or cutting the unstable hinge between domains.[148]
The rationale is that compared to the global, stable and protected region, the other parts of the
protein are more accessible to the protease, so they have a higher possibility for the initial
cleavage.

A few studies combined limited proteolysis and MS for studying protein

41

structure,[149-150] but limited proteolysis usually serves as a supplement to traditional x-ray
crystallography and two-dimensional NMR analysis.
Burley’s group recently employs a high-throughput protocol for protein domain
elucidation. They apply 96-well arrays for limited digestion in orthogonal conditions (four
proteases and six reaction times). From an experiment with 400 target proteins, they draw
statistical conclusions, most notably that proteins digested to contain only stable globular
domains have the highest probability for crystallization (and subsequent structure determination).
In conclusion, they believe the high throughput limited digestion process can effectively gather
the domain information for the protein and determine the optimal construct selection to facilitate
crystallization and protein structure determination.[151]
Sharing the same concept that stable and globular proteins have a high tendency to
crystallize, Edwards and coworkers add trace amounts of protease (enzyme/substrate ratio from
1:10000 to 1:100 wt/wt) into protein crystallization solutions, and find that in situ limited
proteolysis promotes the crystallization of 55 bacterial and 14 human proteins.[152] Although
the crystals are sometimes difficult to reproduce, the idea of using limited proteolysis to improve
crystallization is worth considering along with systematic methods for limited digestion.
Yamaguchi, Miyazaki and Maeda mention limited proteolysis with respect to proteomics.
[153] They integrate protease-containing microreactors into the digestion prior to MS analysis,
and decrease the in-solution reaction from 16-h to a few minutes. However, the “limited
proteolysis” in their study refers more to rapid and efficient digestion than its traditional
definition.
The innovative and excellent work described above serves as an inspiration to utilize the
unique property of protease-containing membranes for limited digestion in proteomics research.

42

The dissertation (chapter 4 and 5) describes limited proteolysis via msec-digestion in a proteasecontaining membrane and the potential application of this technique in middle-down proteomics
and low resolution probing of protein structure.

1.6

Outline of Dissertation
The following dissertation contains five chapters. The basic structure is a fundamental

study followed by a chapter with one or more biological applications. Thus, chapters 2 and 3
describe the development and characterization of TiO2-modified membranes, and an
investigation primarily of tau phosphorylation, respectively. Similarly, chapter 4 discusses the
limited proteolysis of model proteins, bovine serum albumin and apomyoglobin, in a proteasemodified membrane, and the subsequent chapter explores limited proteolysis for probing the
structure of a root hair defective protein. Finally, chapter 6 summarizes the above work and
provides some future research directions. Specifically, the next five chapter titles are:
Chapter 2: Phosphopeptide Enrichment with TiO2-Modified Membranes
Chapter 3: Investigation of Phosphorylation in Tau Protein and the Nuclear Factor Kappa B
(NF-B) Pathway using Phosphopeptide Enrichment with TiO2-modified Membranes
Chapter 4: Limited Proteolysis Via msec-Digestions in Protease-Modified Membranes
Chapter 5: Protease-Modified Membranes for Studies of Protein Structure: Limited Proteolysis
and Rapid Digestion for HDX
Chapter 6: Summary and Future Work

43

REFERENCES

44

1.7

References

1.

Yarmush, M. L.; Jayaraman, A., Advances in Proteomic Technologies. Annu. Rev.
Biomed. Eng. 2002, 4 (1), 349-373.

2.

Yates, J. R.; Ruse, C. I.; Nakorchevsky, A., Proteomics by Mass Spectrometry:
Approaches, Advances, and Applications. Annu. Rev. Biomed. Eng. 2009, 11 (1), 49-79.

3.

Han, X.; Aslanian, A.; Yates Iii, J. R., Mass Spectrometry for Proteomics. Curr. Opin.
Chem. Biol. 2008, 12 (5), 483-490.

4.

Karas, M.; Bachmann, D.; Hillenkamp, F., Influence of the Wavelength in HighIrradiance Ultraviolet Laser Desorption Mass Spectrometry of Organic Molecules. Anal.
Chem. 1985, 57 (14), 2935-2939.

5.

Yamashita, M.; Fenn, J. B., Electrospray Ion Source. Another Variation on the Free-Jet
Theme. J. Phys. Chem. 1984, 88 (20), 4451-4459.

6.

Aebersold, R.; Mann, M., Mass Spectrometry-Based Proteomics. Nature 2003, 422
(6928), 198-207.

7.

Domon, B.; Aebersold, R., Mass Spectrometry and Protein Analysis. Science 2006, 312
(5771), 212-217.

8.

Tanaka, K., The Origin of Macromolecule Ionization by Laser Irradiation (Nobel
Lecture). Angew. Chem. Int. Ed. 2003, 42 (33), 3860-3870.

9.

Liao, P.-C.; Allison, J., Dissecting Matrix-Assisted Laser Desorption/Ionization Mass
Spectra. J. Mass Spectrom. 1995, 30 (5), 763-766.

10.

Taylor, G., Disintegration of Water Drops in an Electric Field. Proc. R. Soc. Lond. A,
Math. Phys. Sci. 1964, 280 (1382), 383-397.

11.

Felitsyn, N.; Peschke, M.; Kebarle, P., Origin and Number of Charges Observed on
Multiply-Protonated Native Proteins Produced by ESI. Int. J. Mass spectrom. 2002, 219
(1), 39-62.
45

12.

Martin, S. E.; Shabanowitz, J.; Hunt, D. F.; Marto, J. A., Subfemtomole MS and MS/MS
Peptide Sequence Analysis Using Nano-HPLC Micro-ESI Fourier Transform Ion
Cyclotron Resonance Mass Spectrometry. Anal. Chem. 2000, 72 (18), 4266-4274.

13.

Taká Z.; Wiseman, J. M.; Gologan, B.; Cooks, R. G., Mass Spectrometry Sampling
ts,
under Ambient Conditions with Desorption Electrospray Ionization. Science 2004, 306
(5695), 471-473.

14.

Cody, R. B.; Laramé J. A.; Durst, H. D., Versatile New Ion Source for the Analysis of
e,
Materials in Open Air under Ambient Conditions. Anal. Chem. 2005, 77 (8), 2297-2302.

15.

Douglas, D. J.; Frank, A. J.; Mao, D., Linear Ion Traps in Mass Spectrometry. Mass
Spectrom. Rev. 2005, 24 (1), 1-29.

16.

Papayannopoulos, I. A., The Interpretation of Collision-Induced Dissociation Tandem
Mass Spectra of Peptides. Mass Spectrom. Rev. 1995, 14 (1), 49-73.

17.

Makarov, A., Electrostatic Axially Harmonic Orbital Trapping:  A High-Performance
Technique of Mass Analysis. Anal. Chem. 2000, 72 (6), 1156-1162.

18.

Hu, Q.; Noll, R. J.; Li, H.; Makarov, A.; Hardman, M.; Graham Cooks, R., The Orbitrap:
A New Mass Spectrometer. J. Mass Spectrom. 2005, 40 (4), 430-443.

19.

Roepstorff, P.; Fohlman, J., Letter to the Editors. Biol. Mass Spectrom. 1984, 11 (11),
601-601.

20.

Kelleher, N. L.; Lin, H. Y.; Valaskovic, G. A.; Aaserud, D. J.; Fridriksson, E. K.;
McLafferty, F. W., Top Down Versus Bottom up Protein Characterization by Tandem
High-Resolution Mass Spectrometry. J. Am. Chem. Soc. 1999, 121 (4), 806-812.

21.

Eng, J.; McCormack, A.; Yates, J., An Approach to Correlate Tandem Mass Spectral
Data of Peptides with Amino Acid Sequences in a Protein Database. J. Am. Soc. Mass
Spectrom. 1994, 5 (11), 976-989.

22.

Reid, G. E.; McLuckey, S. A., ‘Top Down’ Protein Characterization Via Tandem Mass
Spectrometry. J. Mass Spectrom. 2002, 37 (7), 663-675.

46

23.

McLafferty, F. W.; Breuker, K.; Jin, M.; Han, X.; Infusini, G.; Jiang, H.; Kong, X.;
Begley, T. P., Top-Down MS, a Powerful Complement to the High Capabilities of
Proteolysis Proteomics. FEBS J. 2007, 274 (24), 6256-6268.

24.

Kellie, J. F.; Tran, J. C.; Lee, J. E.; Ahlf, D. R.; Thomas, H. M.; Ntai, I.; Catherman, A.
D.; Durbin, K. R.; Zamdborg, L.; Vellaichamy, A.; Thomas, P. M.; Kelleher, N. L., The
Emerging Process of Top Down Mass Spectrometry for Protein Analysis: Biomarkers,
Protein-Therapeutics, and Achieving High Throughput. Mol. Biosyst. 2010, 6 (9), 15321539.

25.

Liu,
J.;
Huang,
T.-Y.;
McLuckey,
S.
A.,
Top-Down
Protein
Identification/Characterization of a Priori Unknown Proteins Via Ion Trap CollisionInduced Dissociation and Ion/Ion Reactions in a Quadrupole/Time-of-Flight Tandem
Mass Spectrometer. Anal. Chem. 2009, 81 (4), 1433-1441.

26.

Zubarev, R. A.; Kelleher, N. L.; McLafferty, F. W., Electron Capture Dissociation of
Multiply Charged Protein Cations. A Nonergodic Process. J. Am. Chem. Soc. 1998, 120
(13), 3265-3266.

27.

Syka, J. E. P.; Coon, J. J.; Schroeder, M. J.; Shabanowitz, J.; Hunt, D. F., Peptide and
Protein Sequence Analysis by Electron Transfer Dissociation Mass Spectrometry. Proc.
Natl. Acad. Sci. U. S. A. 2004, 101 (26), 9528-9533.

28.

Siuti, N.; Kelleher, N. L., Decoding Protein Modifications Using Top-Down Mass
Spectrometry. Nat. Methods 2007, 4 (10), 817-821.

29.

Armirotti, A.; Damonte, G., Achievements and Perspectives of Top-Down Proteomics.
Proteomics 2010, 10 (20), 3566-3576.

30.

Garcia, B. A.; Siuti, N.; Thomas, C. E.; Mizzen, C. A.; Kelleher, N. L., Characterization
of Neurohistone Variants and Post-Translational Modifications by Electron Capture
Dissociation Mass Spectrometry. Int. J. Mass spectrom. 2007, 259 (1–3), 184-196.

31.

Cannon, J.; Lohnes, K.; Wynne, C.; Wang, Y.; Edwards, N.; Fenselau, C., HighThroughput Middle-Down Analysis Using an Orbitrap. J. Proteome Res. 2010, 9 (8),
3886-3890.

32.

Garcia, B. A.; Shabanowitz, J.; Hunt, D. F., Characterization of Histones and Their PostTranslational Modifications by Mass Spectrometry. Curr. Opin. Chem. Biol. 2007, 11 (1),
66-73.
47

33.

Lee, M.; Goodbourn, S., Signalling from the Cell Surface to the Nucleus. Regulation of
Gene Expression 2001, 37, 71-85.

34.

Ficarro, S. B.; McCleland, M. L.; Stukenberg, P. T.; Burke, D. J.; Ross, M. M.;
Shabanowitz, J.; Hunt, D. F.; White, F. M., Phosphoproteome Analysis by Mass
Spectrometry and Its Application to Saccharomyces Cerevisiae. Nat. Biotechnol. 2002,
20 (3), 301-305.

35.

Manning, G.; Whyte, D. B.; Martinez, R.; Hunter, T.; Sudarsanam, S., The Protein
Kinase Complement of the Human Genome. Science 2002, 298 (5600), 1912-1934.

36.

Hunter, T., Signaling - 2000 and Beyond. Cell 2000, 100 (1), 113-127.

37.

Graves, J. D.; Krebs, E. G., Protein Phosphorylation and Signal Transduction. Pharmacol.
Ther. 1999, 82 (2-3), 111-121.

38.

Mann, M.; Ong, S. E.; Gronborg, M.; Steen, H.; Jensen, O. N.; Pandey, A., Analysis of
Protein Phosphorylation Using Mass Spectrometry: Deciphering the Phosphoproteome.
Trends Biotechnol. 2002, 20 (6), 261-268.

39.

Olsen, J. V.; Blagoev, B.; Gnad, F.; Macek, B.; Kumar, C.; Mortensen, P.; Mann, M.,
Global, in vivo, and Site-Specific Phosphorylation Dynamics in Signaling Networks. Cell
2006, 127 (3), 635-648.

40.

Johnson, L.; Lewis, R., Structural Basis for Control by Phosphorylation. Chem. Rev. 2001,
101, 2209-2242.

41.

Cohen, P., The Regulation of Protein Function by Multisite Phosphorylation - a 25 Year
Update. Trends Biochem. Sci 2000, 25 (12), 596-601.

42.

Thingholm, T. E.; Jensen, O. N.; Larsen, M. R., Analytical Strategies for
Phosphoproteomics. Proteomics 2009, 9 (6), 1451-1468.

43.

Walsh, C. T., Posttranslational Modification of Proteins: Expending Nature's Inventory.
Roberts and Company Publishers: Englewood, Colorado, USA, 2006.

44.

Cohen, P., The Origins of Protein Phosphorylation. Nat. Cell Biol. 2002, 4 (5), E127E130.
48

45.

Towbin, H.; Staehelin, T.; Gordon, J., Electrophoretic Transfer of Proteins from
Polyacrylamide Gels to Nitrocellulose Sheets: Procedure and Some Applications. Proc.
Natl. Acad. Sci. U. S. A. 1979, 76 (9), 4350-4354.

46.

Zhang, H.; Zha, X.; Tan, Y.; Hornbeck, P. V.; Mastrangelo, A. J.; Alessi, D. R.;
Polakiewicz, R. D.; Comb, M. J., Phosphoprotein Analysis Using Antibodies Broadly
Reactive against Phosphorylated Motifs. J. Biol. Chem. 2002, 277 (42), 39379-39387.

47.

Izaguirre, G.; Aguirre, L.; Ji, P.; Aneskievich, B.; Haimovich, B., Tyrosine
Phosphorylation of Α-Actinin in Activated Platelets. J. Biol. Chem. 1999, 274 (52),
37012-37020.

48.

Zhou, H.; Watts, J. D.; Aebersold, R., A Systematic Approach to the Analysis of Protein
Phosphorylation. Nat. Biotechnol. 2001, 19 (4), 375-378.

49.

Kalume, D. E.; Molina, H.; Pandey, A., Tackling the Phosphoproteome: Tools and
Strategies. Curr. Opin. Chem. Biol. 2003, 7 (1), 64-69.

50.

Craig, A. G.; Hoeger, C. A.; Miller, C. L.; Goedken, T.; Rivier, J. E.; Fischer, W. H.,
Monitoring Protein-Kinase and Phosphatase Reactions with Matrix-Assisted LaserDesorption Ionization Mass-Spectrometry and Capillary Zone Electrophoresis Comparison of the Detection Efficiency of Peptide-Phosphopeptide Mixtures. Biol. Mass
Spectrom. 1994, 23 (8), 519-528.

51.

Liao, P. C.; Leykam, J.; Andrews, P. C.; Gage, D. A.; Allison, J., An Approach to Locate
Phosphorylation Sites in a Phosphoprotein - Mass Mapping by Combining Specific
Enzymatic Degradation with Matrix-Assisted Laser-Desorption Ionization MassSpectrometry. Anal. Biochem. 1994, 219 (1), 9-20.

52.

Steen, H.; Jebanathirajah, J. A.; Rush, J.; Morrice, N.; Kirschner, M. W., Phosphorylation
Analysis by Mass Spectrometry. Mol. Cell. Proteomics 2006, 5 (1), 172-181.

53.

Krause, E.; Wenschuh, H.; Jungblut, P. R., The Dominance of Arginine-Containing
Peptides in MALDI-Derived Tryptic Mass Fingerprints of Proteins. Anal. Chem. 1999,
71 (19), 4160-4165.

54.

Clipston, N. L.; Jai-nhuknan, J.; Cassady, C. J., A Comparison of Negative and Positive
Ion Time-of-Flight Post-Source Decay Mass Spectrometry for Peptides Containing Basic
Residues. Int. J. Mass spectrom. 2003, 222 (1), 363-381.
49

55.

Asara, J. M.; Allison, J., Enhanced Detection of Phosphopeptides in Matrix-Assisted
Laser Desorption/Ionization Mass Spectrometry Using Ammonium Salts. J. Am. Soc.
Mass Spectrom. 1999, 10 (1), 35-44.

56.

Kjellstrom, S.; Jensen, O. N., Phosphoric Acid as a Matrix Additive for MALDI MS
Analysis of Phosphopeptides and Phosphoproteins. Anal. Chem. 2004, 76 (17), 51095117.

57.

Paradela, A.; Albar, J. P., Advances in the Analysis of Protein Phosphorylation. J.
Proteome Res. 2008, 7 (5), 1809-1818.

58.

Reinders, J.; Sickmann, A., State-of-the-Art in Phosphoproteomics. Proteomics 2005, 5
(16), 4052-4061.

59.

Pandey, A.; Fernandez, M. M.; Steen, H.; Blagoev, B.; Nielsen, M. M.; Roche, S.; Mann,
M.; Lodish, H. F., Identification of a Novel Immunoreceptor Tyrosine-Based Activation
Motif-Containing Molecule, Stam2, by Mass Spectrometry and Its Involvement in
Growth Factor and Cytokine Receptor Signaling Pathways. J. Biol. Chem. 2000, 275 (49),
38633-38639.

60.

Steen, H.; Kuster, B.; Fernandez, M.; Pandey, A.; Mann, M., Tyrosine Phosphorylation
Mapping of the Epidermal Growth Factor Receptor Signaling Pathway. J. Biol. Chem.
2002, 277 (2), 1031-1039.

61.

Collins, M. O.; Yu, L.; Choudhary, J. S., Analysis of Protein Phosphorylation on a
Proteome-Scale. Proteomics 2007, 7 (16), 2751-2768.

62.

Gronborg, M.; Kristiansen, T. Z.; Stensballe, A.; Andersen, J. S.; Ohara, O.; Mann, M.;
Jensen, O. N.; Pandey, A., A Mass Spectrometry-Based Proteomic Approach for
Identification of Serine/Threonine-Phosphorylated Proteins by Enrichment with PhosphoSpecific Antibodies - Identification of a Novel Protein, Frigg, as a Protein Kinase a
Substrate. Mol. Cell. Proteomics 2002, 1 (7), 517-527.

63.

Oda, Y.; Nagasu, T.; Chait, B. T., Enrichment Analysis of Phosphorylated Proteins as a
Tool for Probing the Phosphoproteome. Nat. Biotechnol. 2001, 19 (4), 379-382.

64.

Zhou, H. L.; Watts, J. D.; Aebersold, R., A Systematic Approach to the Analysis of
Protein Phosphorylation. Nat. Biotechnol. 2001, 19 (4), 375-378.

50

65.

McLachlin, D. T.; Chait, B. T., Improved Beta-Elimination-Based Affinity Purification
Strategy for Enrichment of Phosphopeptides. Anal. Chem. 2003, 75 (24), 6826-6836.

66.

Porath, J.; Carlsson, J. A. N.; Olsson, I.; Belfrage, G., Metal Chelate Affinity
Chromatography, a New Approach to Protein Fractionation. Nature 1975, 258 (5536),
598-599.

67.

Hochuli, E.; Dö
beli, H.; Schacher, A., New Metal Chelate Adsorbent Selective for
Proteins and Peptides Containing Neighbouring Histidine Residues. J. Chromatogr. 1987,
411 (0), 177-184.

68.

Andersson, L.; Porath, J., Isolation of Phosphoproteins by Immobilized Metal (Fe-3+)
Affinity-Chromatography. Anal. Biochem. 1986, 154 (1), 250-254.

69.

Han, G. H.; Ye, M. L.; Zou, H. F., Development of Phosphopeptide Enrichment
Techniques for Phosphoproteome Analysis. Analyst 2008, 133 (9), 1128-1138.

70.

Feng, S.; Pan, C. S.; Jiang, X. G.; Xu, S. Y.; Zhou, H. J.; Ye, M. L.; Zou, H. F., Fe3+
Immobilized Metal Affinity Chromatography with Silica Monolithic Capillary Column
for Phosphoproteome Analysis. Proteomics 2007, 7 (3), 351-360.

71.

Xu, X. Q.; Deng, C. H.; Gao, M. X.; Yu, W. J.; Yang, P. Y.; Zhang, X. M., Synthesis of
Magnetic Microspheres with Immobilized Metal Ions for Enrichment and Direct
Determination of Phosphopeptides by Matrix-Assisted Laser Desorption Ionization Mass
Spectrometry. Adv. Mater. 2006, 18 (24), 3289-+.

72.

Dunn, J. D.; Igrisan, E. A.; Palumbo, A. M.; Reid, G. E.; Bruening, M. L.,
Phosphopeptide Enrichment Using MALDI Plates Modified with High-Capacity Polymer
Brushes. Anal. Chem. 2008, 80 (15), 5727-5735.

73.

Ikeguchi, Y.; Nakamura, H., Determination of Organic Phosphates by Column-Switching
High Performance Anion-Exchange Chromatography Using on-Line Preconcentration on
Titania. Anal. Sci. 1997, 13 (3), 479-483.

74.

Ikeguchi, Y.; Nakamura, H., Selective Enrichment of Phospholipids by Titania. Anal. Sci.
2000, 16 (5), 541-543.

51

75.

Kawahara, M.; Nakamura, H.; Nakajima, T., Group Separation of Ribonucleosides and
Deoxyribonucleosides on a New Ceramic Titania Column. Anal. Sci. 1989, 5 (6), 763764.

76.

Matsuda, H.; Nakamura, H.; Nakajima, T., New Ceramic Titania - Selective Adsorbent
for Organic-Phosphates. Anal. Sci. 1990, 6 (6), 911-912.

77.

Kosmulski, M., The Significance of the Difference in the Point of Zero Charge between
Rutile and Anatase. Adv. Colloid Interface Sci. 2002, 99 (3), 255-264.

78.

Connor, P. A.; McQuillan, A. J., Phosphate Adsorption onto TiO2 from Aqueous
Solutions: An in Situ Internal Reflection Infrared Spectroscopic Study. Langmuir 1999,
15 (8), 2916-2921.

79.

Larsen, M. R.; Thingholm, T. E.; Jensen, O. N.; Roepstorff, P.; Jorgensen, T. J. D.,
Highly Selective Enrichment of Phosphorylated Peptides from Peptide Mixtures Using
Titanium Dioxide Microcolumns. Mol. Cell. Proteomics 2005, 4 (7), 873-886.

80.

Pinkse, M. W. H.; Uitto, P. M.; Hilhorst, M. J.; Ooms, B.; Heck, A. J. R., Selective
Isolation at the Femtomole Level of Phosphopeptides from Proteolytic Digests Using 2dNanoLC-ESI-MS/MS and Titanium Oxide Precolumns. Anal. Chem. 2004, 76 (14),
3935-3943.

81.

Jensen, S. S.; Larsen, M. R., Evaluation of the Impact of Some Experimental Procedures
on Different Phosphopeptide Enrichment Techniques. Rapid Commun. Mass Spectrom.
2007, 21 (22), 3635-3645.

82.

Thingholm, T. E.; Jorgensen, T. J. D.; Jensen, O. N.; Larsen, M. R., Highly Selective
Enrichment of Phosphorylated Peptides Using Titanium Dioxide. Nat. Protoc. 2006, 1 (4),
1929-1935.

83.

Hata, K.; Morisaka, H.; Hara, K.; Mima, J.; Yumoto, N.; Tatsu, Y.; Furuno, M.; Ishizuka,
N.; Ueda, M., Two-Dimensional HPLC on-Line Analysis of Phosphopeptides Using
Titania and Monolithic Columns. Anal. Biochem. 2006, 350 (2), 292-297.

84.

Li, Y.; Xu, X. Q.; Qi, D. W.; Deng, C. H.; Yang, P. Y.; Zhang, X. M., Novel
Fe3O4@TiO2 Core-Shell Microspheres for Selective Enrichment of Phosphopeptides in
Phosphoproteome Analysis. J. Proteome Res. 2008, 7 (6), 2526-2538.

52

85.

Pinkse, M. W. H.; Mohammed, S.; Gouw, L. W.; van Breukelen, B.; Vos, H. R.; Heck, A.
J. R., Highly Robust, Automated, and Sensitive on Line TiO2-Based Phosphoproteomics
Applied to Study Endogenous Phosphorylation in Drosophila Melanogaster. J. Proteome
Res. 2008, 7 (2), 687-697.

86.

Thingholm, T. E.; Larsen, M. R.; Ingrell, C. R.; Kassem, M.; Jensen, O. N., TiO2-Based
Phosphoproteomic Analysis of the Plasma Membrane and the Effects of Phosphatase
Inhibitor Treatment. J. Proteome Res. 2008, 7 (8), 3304-3313.

87.

Yu, L. R.; Zhu, Z. Y.; Chan, K. C.; Issaq, H. J.; Dimitrov, D. S.; Veenstra, T. D.,
Improved Titanium Dioxide Enrichment of Phosphopeptides from Hela Cells and High
Confident Phosphopeptide Identification by Cross-Validation of MS/MS and MS/MS/MS
Spectra. J. Proteome Res. 2007, 6 (11), 4150-4162.

88.

Rainer, M.; Sonderegger, H.; Bakry, R.; Huck, C. W.; Morandell, S.; Huber, L. A.;
Gjerde, D. T.; Bonn, G. K., Analysis of Protein Phosphorylation by Monolithic
Extraction Columns Based on Poly(Divinylbenzene) Containing Embedded Titanium
Dioxide and Zirconium Dioxide Nano-Powders. Proteomics 2008, 8 (21), 4593-4602.

89.

Cantin, G. T.; Shock, T. R.; Park, S. K.; Madhani, H. D.; Yates, J. R., Optimizing TiO2Based Phosphopeptide Enrichment for Automated Multidimensional Liquid
Chromatography Coupled to Tandem Mass Spectrometry. Anal. Chem. 2007, 79 (12),
4666-4673.

90.

Trinidad, J. C.; Specht, C. G.; Thalhammer, A.; Schoepfer, R.; Burlingame, A. L.,
Comprehensive Identification of Phosphorylation Sites in Postsynaptic Density
Preparations. Mol. Cell. Proteomics 2006, 5 (5), 914-922.

91.

Sugiyama, N.; Masuda, T.; Shinoda, K.; Nakamura, A.; Tomita, M.; Ishihama, Y.,
Phosphopeptide Enrichment by Aliphatic Hydroxy Acid-Modified Metal Oxide
Chromatography for Nano-LC-MS/MS in Proteomics Applications. Mol. Cell.
Proteomics 2007, 6 (6), 1103-1109.

92.

Yu, L.-R.; Zhu, Z.; Chan, K. C.; Issaq, H. J.; Dimitrov, D. S.; Veenstra, T. D., Improved
Titanium Dioxide Enrichment of Phosphopeptides from Hela Cells and High Confident
Phosphopeptide Identification by Cross-Validation of MS/MS and MS/MS/MS Spectra. J.
Proteome Res. 2007, 6 (11), 4150-4162.

93.

Simon, E. S.; Young, M.; Chan, A.; Bao, Z. Q.; Andrews, P. C., Improved Enrichment
Strategies for Phosphorylated Peptides on Titanium Dioxide Using Methyl Esterification
and pH Gradient Elution. Anal. Biochem. 2008, 377 (2), 234-242.
53

94.

Tsai, C. F.; Wang, Y. T.; Chen, Y. R.; Lai, C. Y.; Lin, P. Y.; Pan, K. T.; Chen, J. Y.;
Khoo, K. H.; Chen, Y. J., Immobilized Metal Affinity Chromatography Revisited:
pH/Acid Control toward High Selectivity in Phosphoproteomics. J. Proteome Res. 2008,
7 (9), 4058-4069.

95.

Thingholm, T. E.; Jensen, O. N.; Robinson, P. J.; Larsen, M. R., Simac (Sequential
Elution from IMAC), a Phosphoproteomics Strategy for the Rapid Separation of
Monophosphorylated from Multiply Phosphorylated Peptides. Mol. Cell. Proteomics
2008, 7 (4), 661-671.

96.

Bodenmiller, B.; Mueller, L. N.; Mueller, M.; Domon, B.; Aebersold, R., Reproducible
Isolation of Distinct, Overlapping Segments of the Phosphoproteome. Nat. Methods 2007,
4 (3), 231-237.

97.

Liang, S. S.; Makamba, H.; Huang, S. Y.; Chen, S. H., Nano-Titanium Dioxide
Composites for the Enrichment of Phosphopeptides. J. Chromatogr. 2006, 1116 (1-2),
38-45.

98.

Villé J.; Gygi, S. P., The Scx/IMAC Enrichment Approach for Global Phosphorylation
n,
Analysis by Mass Spectrometry. Nat. Protoc. 2008, 3 (10), 1630-1638.

99.

Villé J.; Beausoleil, S. A.; Gerber, S. A.; Gygi, S. P., Large-Scale Phosphorylation
n,
Analysis of Mouse Liver. Proc. Natl. Acad. Sci. U. S. A. 2007, 104 (5), 1488-1493.

100.

Zhai, B.; Villé J.; Beausoleil, S. A.; Mintseris, J.; Gygi, S. P., Phosphoproteome
n,
Analysis of Drosophila Melanogaster Embryos. J. Proteome Res. 2008, 7 (4), 1675-1682.

101.

Palumbo, A. M.; Smith, S. A.; Kalcic, C. L.; Dantus, M.; Stemmer, P. M.; Reid, G. E.,
Tandem Mass Spectrometry Strategies for Phosphoproteome Analysis. Mass Spectrom.
Rev. 2011, 30 (4), 600-625.

102.

Palumbo, A. M.; Reid, G. E., Evaluation of Gas-Phase Rearrangement and Competing
Fragmentation Reactions on Protein Phosphorylation Site Assignment Using Collision
Induced Dissociation-MS/MS and MS(3). Anal. Chem. 2008, 80 (24), 9735-9747.

103.

Olsen, J. V.; Macek, B.; Lange, O.; Makarov, A.; Horning, S.; Mann, M., Higher-Energy
C-Trap Dissociation for Peptide Modification Analysis. Nat. Methods 2007, 4 (9), 709712.

54

104.

Coon, J. J., Collisions or Electrons? Protein Sequence Analysis in the 21st Century. Anal.
Chem. 2009, 81 (9), 3208-3215.

105.

Swaney, D. L.; Wenger, C. D.; Thomson, J. A.; Coon, J. J., Human Embryonic Stem Cell
Phosphoproteome Revealed by Electron Transfer Dissociation Tandem Mass
Spectrometry. Proc. Natl. Acad. Sci. U. S. A. 2009, 106 (4), 995-1000.

106.

Good, D. M.; Wirtala, M.; McAlister, G. C.; Coon, J. J., Performance Characteristics of
Electron Transfer Dissociation Mass Spectrometry. Mol. Cell. Proteomics 2007, 6 (11),
1942-1951.

107.

Nelson, J. M.; Griffin, E. G., Adsorption of Invertase. J. Am. Chem. Soc. 1916, 38 (5),
1109-1115.

108.

Ma, J.; Zhang, L.; Liang, Z.; Shan, Y.; Zhang, Y., Immobilized Enzyme Reactors in
Proteomics. TrAC, Trends Anal. Chem. 2011, 30 (5), 691-702.

109.

Girelli, A. M.; Mattei, E., Application of Immobilized Enzyme Reactor in on-Line High
Performance Liquid Chromatography: A Review. J. Chromatogr. B 2005, 819 (1), 3-16.

110.

Kim, J.; Kim, B. C.; Lopez-Ferrer, D.; Petritis, K.; Smith, R. D., Nanobiocatalysis for
Protein Digestion in Proteomic Analysis. Proteomics 2010, 10 (4), 687-699.

111.

Krenkova, J.; Svec, F., Less Common Applications of Monoliths: Iv. Recent
Developments in Immobilized Enzyme Reactors for Proteomics and Biotechnology. J.
Sep. Sci. 2009, 32 (5-6), 706-718.

112.

Cooper, J. W.; Chen, J.; Li, Y.; Lee, C. S., Membrane-Based Nanoscale Proteolytic
Reactor Enabling Protein Digestion, Peptide Separation, and Protein Identification Using
Mass Spectrometry. Anal. Chem. 2003, 75 (5), 1067-1074.

113.

Smuleac, V.; Butterfield, D. A.; Bhattacharyya, D., Layer-by-Layer-Assembled
Microfiltration Membranes for Biomolecule Immobilization and Enzymatic Catalysis.
Langmuir 2006, 22 (24), 10118-10124.

114.

Xu, F.; Wang, W. H.; Tan, Y. J.; Bruening, M. L., Facile Trypsin Immobilization in
Polymeric Membranes for Rapid, Efficient Protein Digestion. Anal. Chem. 2010, 82 (24),
10045-10051.

55

115.

Liu, Y.; Zhong, W.; Meng, S.; Kong, J.; Lu, H.; Yang, P.; Girault, H. H.; Liu, B.,
Assembly-Controlled Biocompatible Interface on a Microchip: Strategy to Highly
Efficient Proteolysis. Chemistry – A European Journal 2006, 12 (25), 6585-6591.

116.

Bhattacharjee, S.; Dong, J.; Ma, Y.; Hovde, S.; Geiger, J. H.; Baker, G. L.; Bruening, M.
L., Formation of High-Capacity Protein-Adsorbing Membranes through Simple
Adsorption of Poly(Acrylic Acid)-Containing Films at Low pH. Langmuir 2012, 28 (17),
6885-6892.

117.

Liu, G.; Dotzauer, D. M.; Bruening, M. L., Ion-Exchange Membranes Prepared Using
Layer-by-Layer Polyelectrolyte Deposition. J. Membr. Sci. 2010, 354 (1–2), 198-205.

118.

Sproß J.; Sinz, A., A Capillary Monolithic Trypsin Reactor for Efficient Protein
,
Digestion in Online and Offline Coupling to ESI and MALDI Mass Spectrometry. Anal.
Chem. 2010, 82 (4), 1434-1443.

119.

Calleri, E.; Temporini, C.; Furlanetto, S.; Loiodice, F.; Fracchiolla, G.; Massolini, G.,
Lipases for Biocatalysis: Development of a Chromatographic Bioreactor. J. Pharm.
Biomed. Anal. 2003, 32 (4–5), 715-724.

120.

Krenkova, J.; Lacher, N. A.; Svec, F., Highly Efficient Enzyme Reactors Containing
Trypsin and Endoproteinase Lysc Immobilized on Porous Polymer Monolith Coupled to
MS Suitable for Analysis of Antibodies. Anal. Chem. 2009, 81 (5), 2004-2012.

121.

Dulay, M. T.; Baca, Q. J.; Zare, R. N., Enhanced Proteolytic Activity of Covalently
Bound Enzymes in Photopolymerized Sol Gel. Anal. Chem. 2005, 77 (14), 4604-4610.

122.

Brady, D.; Jordaan, J., Advances in Enzyme Immobilisation. Biotechnol. Lett 2009, 31
(11), 1639-1650.

123.

Lin, S.; Yao, G.; Qi, D.; Li, Y.; Deng, C.; Yang, P.; Zhang, X., Fast and Efficient
Proteolysis by Microwave-Assisted Protein Digestion Using Trypsin-Immobilized
Magnetic Silica Microspheres. Anal. Chem. 2008, 80 (10), 3655-3665.

124.

Lin, S.; Yun, D.; Qi, D.; Deng, C.; Li, Y.; Zhang, X., Novel Microwave-Assisted
Digestion by Trypsin-Immobilized Magnetic Nanoparticles for Proteomic Analysis. J.
Proteome Res. 2008, 7 (3), 1297-1307.

56

125.

Xu, F.; Zhen, G.; Yu, F.; Kuennemann, E.; Textor, M.; Knoll, W., Combined Affinity
and Catalytic Biosensor:  In Situ Enzymatic Activity Monitoring of Surface-Bound
Enzymes. J. Am. Chem. Soc. 2005, 127 (38), 13084-13085.

126.

Bruening, M. L.; Dotzauer, D. M.; Jain, P.; Ouyang, L.; Baker, G. L., Creation of
Functional Membranes Using Polyelectrolyte Multilayers and Polymer Brushes.
Langmuir 2008, 24 (15), 7663-7673.

127.

Lee, T. D.; Shively, J. E., [19] Enzymatic and Chemical Digestion of Proteins for Mass
Spectrometry. In Methods Enzymol., James, A. M., Ed. Academic Press: 1990; Vol.
Volume 193, pp 361-374.

128.

Qian, K.; Wan, J.; Qiao, L.; Huang, X.; Tang, J.; Wang, Y.; Kong, J.; Yang, P.; Yu, C.;
Liu, B., Macroporous Materials as Novel Catalysts for Efficient and Controllable
Proteolysis. Anal. Chem. 2009, 81 (14), 5749-5756.

129.

Bi, H.; Qiao, L.; Busnel, J.-M.; Liu, B.; Girault, H. H., Kinetics of Proteolytic Reactions
in Nanoporous Materials. J. Proteome Res. 2009, 8 (10), 4685-4692.

130.

Gan, J.; Zhu, J.; Yan, G.; Liu, Y.; Yang, P.; Liu, B., Periodic Mesoporous Organosilica as
a Multifunctional Nanodevice for Large-Scale Characterization of Membrane Proteins.
Anal. Chem. 2012, 84 (13), 5809-5815.

131. Nelson, D. L.; Lehninger, A. L.; Cox, M. M., Lehninger Principles of Biochemistry. W.H.
Freeman: New York, 2008.
132.

Hamuro, Y.; Coales, S. J.; Molnar, K. S.; Tuske, S. J.; Morrow, J. A., Specificity of
Immobilized Porcine Pepsin in H/D Exchange Compatible Conditions. Rapid Commun.
Mass Spectrom. 2008, 22 (7), 1041-1046.

133.

Ma, J.; Hou, C.; Sun, L.; Tao, D.; Zhang, Y.; Shan, Y.; Liang, Z.; Zhang, L.; Yang, L.;
Zhang, Y., Coupling Formic Acid Assisted Solubilization and Online Immobilized
Pepsin Digestion with Strong Cation Exchange and Microflow Reversed-Phase Liquid
Chromatography with Electrospray Ionization Tandem Mass Spectrometry for Integral
Membrane Proteome Analysis. Anal. Chem. 2010, 82 (23), 9622-9625.

134.

Liuni, P.; Rob, T.; Wilson, D. J., A Microfluidic Reactor for Rapid, Low-Pressure
Proteolysis with on-Chip Electrospray Ionization. Rapid Commun. Mass Spectrom. 2010,
24 (3), 315-320.
57

135.

Rey, M.; Man, P.; Brandolin, G.; Forest, E.; Pelosi, L., Recombinant Immobilized
Rhizopuspepsin as a New Tool for Protein Digestion in Hydrogen/Deuterium Exchange
Mass Spectrometry. Rapid Commun. Mass Spectrom. 2009, 23 (21), 3431-3438.

136.

Engen, J. R.; Smith, D. L., Peer Reviewed: Investigating Protein Structure and Dynamics
by Hydrogen Exchange MS. Anal. Chem. 2001, 73 (9), 256 A-265 A.

137.

Engen, J. R., Analysis of Protein Conformation and Dynamics by Hydrogen/Deuterium
Exchange MS. Anal. Chem. 2009, 81 (19), 7870-7875.

138.

Englander, S. W.; Mayne, L.; Bai, Y.; Sosnick, T. R., Hydrogen Exchange: The Modern
Legacy of Linderstrø
m-Lang. Protein Sci. 1997, 6 (5), 1101-1109.

139.

Konermann, L.; Pan, J.; Liu, Y.-H., Hydrogen Exchange Mass Spectrometry for Studying
Protein Structure and Dynamics. Chem. Soc. Rev. 2011, 40 (3), 1224-1234.

140.

Katta, V.; Chait, B. T.; Carr, S., Conformational Changes in Proteins Probed by
Hydrogen-Exchange Electrospray-Ionization Mass Spectrometry. Rapid Commun. Mass
Spectrom. 1991, 5 (4), 214-217.

141.

Kaltashov, I. A.; Bobst, C. E.; Abzalimov, R. R., H/D Exchange and Mass Spectrometry
in the Studies of Protein Conformation and Dynamics: Is There a Need for a Top-Down
Approach? Anal. Chem. 2009, 81 (19), 7892-7899.

142.

Pan, J.; Han, J.; Borchers, C. H.; Konermann, L., Hydrogen/Deuterium Exchange Mass
Spectrometry with Top-Down Electron Capture Dissociation for Characterizing
Structural Transitions of a 17 Kda Protein. J. Am. Chem. Soc. 2009, 131 (35), 1280112808.

143.

Eliezer, D.; Yao, J.; Dyson, H. J.; Wright, P. E., Structural and Dynamic Characterization
of Partially Folded States of Apomyoglobin and Implications for Protein Folding. Nat.
Struct. Biol. 1998, 18 (2), 148-155.

144.

Jø
rgensen, T. J. D.; Gå
rdsvoll, H.; Ploug, M.; Roepstorff, P., Intramolecular Migration of
Amide Hydrogens in Protonated Peptides Upon Collisional Activation. J. Am. Chem. Soc.
2005, 127 (8), 2785-2793.

58

145.

Morgan, C. R.; Engen, J. R., Investigating Solution-Phase Protein Structure and
Dynamics by Hydrogen Exchange Mass Spectrometry. In Current Protocols in Protein
Science, John Wiley & Sons, Inc.: 2009.

146.

Rob, T.; Liuni, P.; Gill, P. K.; Zhu, S.; Balachandran, N.; Berti, P. J.; Wilson, D. J.,
Measuring Dynamics in Weakly Structured Regions of Proteins Using MicrofluidicsEnabled Subsecond H/D Exchange Mass Spectrometry. Anal. Chem. 2012, 84 (8), 37713779.

147.

Rey, M.; Man, P.; Clé
menç B.; Tré guet, V.; Brandolin, G.; Forest, E.; Pelosi, L.,
on,
zé
Conformational Dynamics of the Bovine Mitochondrial Adp/Atp Carrier Isoform 1
Revealed by Hydrogen/Deuterium Exchange Coupled to Mass Spectrometry. J. Biol.
Chem. 2010, 285 (45), 34981-34990.

148.

Fontana, A.; Polverino de Laureto, P.; Spolaore, B.; Frare, E.; Zambonin, M., Detecting
Disordered Regions in Proteins by Limited Proteolysis. In Instrumental Analysis of
Intrinsically Disordered Proteins, John Wiley & Sons, Inc.: 2010; pp 569-626.

149.

Cohen, S. L.; Chait, B. T., Mass Spectrometry as a Tool for Protein Crystallography.
Annu. Rev. Biophys. Biomol. Struct. 2001, 30 (1), 67-85.

150.

Cohen, S. L.; Chait, B. T.; Ferré
-D'Amaré A. R.; Burley, S. K., Probing the Solution
,
Structure of the DNA-Binding Protein Max by a Combination of Proteolysis and Mass
Spectrometry. Protein Sci. 1995, 4 (6), 1088-1099.

151.

Gao, X.; Bain, K.; Bonanno, J.; Buchanan, M.; Henderson, D.; Lorimer, D.; Marsh, C.;
Reynes, J.; Sauder, J.; Schwinn, K.; Thai, C.; Burley, S., High-Throughput Limited
Proteolysis/Mass Spectrometry for Protein Domain Elucidation. J. Struct. Funct.
Genomics. 2005, 6 (2), 129-134.

152.

Dong, A.; Xu, X.; Edwards, A. M., In Situ Proteolysis for Protein Crystallization and
Structure Determination. Nat. Methods 2007, 4 (12), 1019-1021.

153.

Yamaguchi, H.; Miyazaki, M.; Maeda, H., Limited Proteolysis in Proteomics Using
Protease-Immobilized Microreactors. Functional Genomics. Kaufmann, M.; Klinger, C.,
Eds. Springer New York: 2012; Vol. 815, pp 187-198.

59

Chapter 2. Phosphopeptide Enrichment with TiO2-Modified
Membranes
(This chapter was submitted for publication as part of an article in Analytical Chemistry)
Selective enrichment of phosphopeptides prior to their analysis by mass spectrometry
(MS) is vital for identifying protein phosphorylation sites involved in cellular regulation. This
chapter describes modification of porous nylon substrates with TiO2 nanoparticles to create
membranes that rapidly enrich phosphopeptides from small-volume samples. Membranes with a
22-mm diameter bind 540 nmol of phosphoangiotensin and recover 70% of the phosphopeptides
in mixtures with a 15-fold excess of unphosphorylated protein digests. Recovery is 90 % for a
pure phosphopeptide. Insertion of small membrane disks into HPLC fittings allows enrichment
of small-volume (10’s of L) samples. Using 2 % TFA as the loading and washing buffer, the
TiO2-modified membranes selectively bind phosphopeptides in a 100-fold excess of
unphosphorylated protein digests.

2.1

Introduction
Phosphorylation of serine, threonine, or tyrosine residues is one of the most important

mechanisms for creating protein diversity in eukaryotic cells.[1] Remarkably, about one-third of
human proteins are substrates for phosphorylation at some stage in their life cycle,[2] and
reversible phosphorylation plays a key role in regulating cell processes such as signal
transduction and metabolism.[3-4] Thus, a fundamental understanding of regulatory pathways
requires identification of protein phosphorylation states. Currently, the leading technology for
evaluating protein phosphorylation is MS,[5] due to its high speed and sensitivity, and the ability
60

in some cases to determine the location of the phosphorylation site by tandem mass spectrometry
(MS/MS) methods.[6] However, although reversible phosphorylation is ubiquitous in cells,
phosphorylation is intrinsically a dynamic and transient property,[7] so phosphoproteins are
generally present at relatively low abundances and low stoichiometries compared to
unphosphorylated proteins.[8] To partially overcome the low abundance, a number of groups
developed methods to enrich phosphopeptides from digested protein mixtures.[9-14]

Most

current phosphoprotein characterization occurs by electrospray ionization (ESI) or matrix
assisted laser desorption/ionization (MALDI) MS following an initial phosphoprotein or
phosphopeptide enrichment procedure.[15-17]
Phosphopeptide

enrichment

methods

include

immunoprecipitation,

covalent

immobilization, and adsorption to metal oxide resins or immobilized metal-ion complexes.[7, 1820]

All of these methods are useful, and they often provide complementary subsets of

phosphopeptides.[21] Studies with adsorption to metal oxides suggest that this technique is
particularly attractive for rapid and highly specific enrichment if binding conditions are carefully
controlled.[17, 22] At low pH, TiO2, the metal oxide most commonly used for phosphopeptide
enrichment, has a positively charged surface that selectively adsorbs phosphorylated species, and
the adsorbed species subsequently elute under basic conditions.[23] Addition of excluders and
low-pH buffer to the phosphopeptide solution helps to alleviate nonspecific binding of acidic
peptides to TiO2.[24-25]
This chapter describes the functionalization of porous nylon substrates with a thin layer
of TiO2 nanoparticles to create membranes for phosphopeptide enrichment.[26] Adsorption of
phosphopeptides from solutions passing through the membrane (Figure 2.1) followed by rinsing
and elution yields concentrated solutions of isolated phosphopeptides. The sub-micron pore size
61

in the nylon allows phosphopeptide adsorption without mass transfer limitations, and the high
surface areas of both the nylon substrate and the nanoparticles give materials with a high
phosphopeptide binding capacity.[25] Moreover, because of the minimal membrane thickness
and relatively large pore size, a syringe is sufficient to push solutions through the membrane
(Figure 2.1).

Figure 2.1. Cartoon of a membrane containing TiO2 for phosphopeptide enrichment and a
miniaturized membrane holder with a syringe pump. PSS=poly(styrene sulfonate).
62

2.2

Experimental

2.2.1 Materials and solutions
Titanium isopropoxide, poly(sodium 4-styrene-sulfonate) (PSS), bovine -casein, bovine
-casein, chicken egg ovalbumin, and bovine serum albumin (BSA) were purchased from
Sigma-Aldrich. BSA was digested using sequencing grade modified porcine trypsin treated with
L-1-tosylamide-2-phenylethyl chloromethyl ketone (Sigma-Aldrich), whereas other proteins
were digested using sequencing grade modified trypsin from Promega. Other reagents employed
in the digestion include Tris-HCl (Invitrogen), urea (J.T.Baker), 1,4-dithio-DL-threitol
(BioChemika), iodoacetamide (Sigma), and ammonium bicarbonate (Columbus Chemical). 2,5dihydroxybenzoic acid (DHB) was purchased from Sigma. Human angiotensin II (DRVYIHPF)
and phosphorylated angiotensin II (DRVpYIHPF) were acquired from Calbiochem.
CH3CH2CO-LFTGHPEpSLEK (H5 peptide) and CD3CD2CO- LFTGHPEpSLEK (D5 peptide)
were synthesized by Dr. Amanda Palumbo from Prof. Gavin E. Reid’s group using manual
stepwise Fmoc-based solid-phase peptide synthesis as described previously.[27]
Enrichment employed loading buffers containing 0.1 vol % trifluoroacetic acid (TFA),
49.9 vol % deionized water and 50 vol % acetonitrile (loading buffer I) or 2 vol % TFA, 48 vol %
deionized water and 50 vol % acetonitrile (loading buffer II). Washing buffer I and II had the
same compositions as loading buffers I and II, respectively. The elution buffer contained 1 vol %
ammonium hydroxide (NH4OH), 49 vol % deionized water and 50 vol % acetonitrile, and had a
pH of 10.9.

63

2.2.2 Membranes and holders
Membrane modification and binding capacity studies employed an Amicon cell (model
2

8010, Millipore) that exposed a membrane area of 3.8 cm . A peristaltic pump pulled solutions
through the membrane. The unmodified nylon membranes (25 mm-diameter, nominal pore size
of 0.45 m) were obtained from Millipore (HNWP02500). For experiments with small sample
volumes (<1 mL), a 6 mm-diameter piece of membrane was placed in a flangeless fitting system
(A-424, Upchurch Scientific) connected to 1/16 inch OD tubing via ferrules. The membrane was
2

supported by a frit in the holder. The membrane area exposed to sample (0.02 cm ) was
determined by passing a dye solution through the apparatus and measuring the diameter of the
colored membrane area with calipers. Sample injection occurred via a syringe pump using a 1mL or 100-L syringe (Figure 2.1).

2.2.3 Synthesis of titania nanoparticles
TiO2 nanoparticles were prepared by the controlled hydrolysis of titanium
isopropoxide.[28] Ti(OCH(CH3)2)4 (1.25 mL) was dissolved in 25 mL of absolute ethanol, and
this solution was added dropwise under vigorous stirring to 250 mL of water previously adjusted
to pH 1.5 with nitric acid. The resulting transparent colloidal suspension was stirred overnight,
and then stored in a refrigerator prior to use.

2.2.4 Membrane modification
Modification of UV/ozone-cleaned (10 min) nylon membranes with titania nanoparticles
occurred through consecutive adsorption of PSS and titania. Ten mL of a 0.02 M PSS, 0.5 M
64

NaCl solution was first passed through the membrane at 2 mL/min, followed by 30 mL of
deionized water at the same flow rate. Subsequently, a 10 mL aliquot of the titania suspension
was circulated through the membrane at 1 mL/min for 30 minutes. Prior to passing through the
membrane, the titania suspension was centrifuged for 5 min at 1300 g to remove large particles.
After TiO2 deposition, the membrane was rinsed with 30 mL of deionized water at 2 mL/min. A
second PSS/TiO2 bilayer was deposited similarly. The amount of TiO2 in the membrane was
determined by inductively coupled plasma-optical emission spectroscopy (ICP-OES) analysis of
TiO2-nanoparticle suspensions before and after passing them through the membrane. A
calibration curve was prepared using a 2 % nitric acid solution containing Ti concentrations from
0.5 to 6 ppm. (Standards were prepared from 998 ppm Ti calibration solution from Sigma) The
suspensions were diluted by a factor of 200 in 2% nitric acid prior to analysis.

2.2.5 Breakthrough curves and determination of binding capacity
In the breakthrough experiments, 50 mL of 20 M phosphoangiotensin in loading buffer
I was passed through a nylon membrane modified with a (PSS/TiO2)2 film, and the effluent was
collected in 2.5-mL aliquots. The flow rate was 2 mL/min. Phosphoangiotensin concentrations in
each aliquot were determined by MALDI-MS using angiotensin as an internal standard. The
amount of bound phosphoangiotensin was determined from the difference in feed and effluent
concentrations in each aliquot.

65

2.2.6 Determination of phosphopeptide recovery using labeled peptides
Ten pmol of H5 peptide (CH3CH2CO-LFTGHPEpSLEK) and 150 pmol of BSA digest
2

were dissolved in 100 L of loading buffer I and passed through the small (0.02 cm ) membrane
at 10 L/min. The membrane was washed with 500 L of washing buffer I at 50 L/min, and
peptides were eluted with 30 L of elution buffer at 3 L/min. One L of eluate (333 fmol of
H5 peptide assuming 100% recovery) and 1 L of internal standard (500 fmol of D5 peptide/L)
were spotted on a MALDI plate followed by 1 L of matrix solution (40 mg/mL of 2,5 DHB in 1%
TFA, 49% H2O, 50% ACN). The ratio of MALDI-MS signals from the H5 and D5 peptides was
compared to a calibration curve to determine the concentration of the H5 peptide in the eluate.

2.2.7 Protein digestion
Bovine -casein, -casein and chicken ovalbumin (100 g each) were first dissolved
separately in 20 L of 6 M urea, 50 mM Tris-HCl. Five L of 10 mM DTT was added to each
protein solution, and the solutions were heated for 1 hour at 56 º to break disulfide bonds.
C
After cooling, 160 L of 50 mM NH4HCO3 and 10 µL of 100 mM iodoacetamide were added to
the solution, which was then allowed to stand for 1 hour in the dark. Finally 10 µL of 0.5 µg/µL
modified trypsin was added to each sample, and the solution was incubated for 16 hours at 37 º
C.
The digestion was quenched by addition of 11 L of acetic acid to lower the pH to about 3. For
the unphosphorylated protein BSA, the digestion followed a similar procedure except the trypsin
employed was sequencing grade modified porcine trypsin from Sigma instead of sequencing
grade trypsin because of the large amount of trypsin required to digest 1 mg of BSA.
66

2.2.8 Phophopeptide enrichment from protein digests
For analysis of phosphopeptides in a large excess of non-phosphorylated peptide, 15
pmol of -casein, -casein, and ovalbumin digests and 1500 pmol of BSA digest were dissolved
in 500 L loading buffer II. This solution was passed through the small-scale TiO2-modified
nylon membrane at 50 L/min, and the membrane was washed with 3 mL of washing buffer II at
100 L/min. Finally the phosphopeptides were eluted from the membrane with 10 L of elution
buffer at 2 L/min .

2.2.9 Mass spectrometry and data analysis
An LTQ XL ion trap mass spectrometer equipped with a vMALDI source (Thermo
Fisher Scientific, San Jose, CA) was used to perform MALDI-MS and CID-MS/MS analyses.
Typically, 1 L of sample solution was spotted onto the stainless steel MALDI plate, followed
by addition of 1 L of matrix solution (40 mg/mL of 2.5 DHB in 1% TFA, 49% H2O, 50% ACN)
and drying. MALDI-MS spectra were acquired using a laser power of 10 µJ across the m/z
range 300-2000 (normal mass range) or 800-4000 (high mass range). CID-MS/MS was
performed on mass-selected precursor ions, using default activation parameters (q=0.25,
activation time 30 msec), with an isolation width of 5 Da, and normalized collision energies from
30-40. Spectra were the average of 60-150 scans. Assignment of peptides resulting from protein
digestion was performed manually by mass fingerprinting or by MS/MS. Phosphorylation sites
were manually assigned by matching MS/MS product ions with in silico peptide fragments
generated using ProteinProspector (http://prospector.ucsf.edu).

67

2.3

Results and discussion

2.3.1 Characterization of TiO2-modified nylon membranes
Nanoparticles are attractive adsorbents for phosphopeptides because of their high surface
area to volume ratio.[29-31] TEM images show that the TiO2 particles in this study have an
average diameter of 5 nm (Figure 2.2), but the particles aggregate into clusters with a diameter of
50 nm. Alternating adsorption of PSS and nanoparticles immobilizes the TiO2 in the membrane,
[26, 31-32] but SEM images of membrane cross-sections do not show the TiO2 nanoparticles
(Figure 2.3c), perhaps because of the rough texture of the nylon membrane and the small and
polydisperse nanoparticle size. However, the Ti energy-dispersive X-ray spectroscopy (EDX)
map in Figure 2.3d reveals the presence of Ti over the whole cross-section of a modified
membrane. Notably, the layer-by-layer method allows modification of nylon membranes with a
PSS/TiO2 film in just 30 min. Moreover, stacking of membranes in the Amicon cell permits
simultaneous modification of several membranes, and a single 25-mm membrane provides 8
membranes for small-scale analysis.

68

a

a

b

b

500 nm

c

100 nm

c
O
Ti

0

1

2

3

4

5

Cu

6

7

8

9

10

Figure 2.2. (a,b) TEM images showing the size distribution of TiO2 nanoparticles at two different
magnifications. (c) EDX analysis of the nanoparticles. The Cu signal arises from the supporting
grid.

69

a

b

5 m

d

c
c

5 m

Figure 2.3. Cross-sectional SEM images of nylon membranes before (a) and after (c) coating
with a PSS/TiO2 nanoparticle film. Images b and d show the EDX Ti maps for the images in (a)
and (c), respectively. In (b) and (d) Ti signals are proportional to the intensity of the red color,
and image b indicates the noise of the measurement.
Nitrogen adsorption isotherms show that a porous nylon membrane (diameter of 25 mm
2

and thickness of 170 m) has a surface area of 3300 cm , which should facilitate the adsorption
of large amounts of TiO2 nanoparticles. Analysis of TiO2 nanoparticle-containing solutions
before and after passing them through the membrane shows that 30 mol of TiO2 adsorb to each

70

membrane (exposed membrane diameter of 22 mm) during deposition of a PSS/TiO2 bilayer
(Figure 2.4). Adsorption of a second PSS/TiO2 bilayer approximately doubles the amount of
TiO2 (55 mol) in the membrane, which should enhance phosphopeptide binding. For 50 nm
8

2

particles, 55 mol corresponds to 7 x 10 particles per cm of internal membrane area, assuming
3

a TiO2 density of 4.2 g/cm .
450000

Intensity (a.u.)

400000

R² 0.9994
=

350000
300000
250000

200000
150000
100000
50000
0
0

1

2

3

4

5

6

7

Concentration / (ppm)
Figure 2.4. Calibration curve for the ICP-OES analysis of Ti. The square and triangle show the
absorbance of the diluted Ti loading solution before and after passing through the nylon
membrane.

2.3.2 Phosphopeptide binding capacity
Binding capacity is important in selectively capturing phosphopeptides because
insufficient capacity may cause low sensitivity or selective detection of only strongly binding

71

phosphopeptides, whereas excess capacity may result in nonspecific binding. We employed
classical breakthrough experiments to determine the amount of phosphopeptide, in this case
phosphoangiotensin, that a (PSS/TiO2)2-modified membrane can capture. As Figure 2.5 shows,
initially the membrane captures all the phosphopeptide in the loading solution, but as the
membrane

becomes

saturated,

phosphopeptides

break

through

and

eventually

no

phosphopeptides bind. Integrating the difference between the feed and permeate phosphopeptide
concentrations

in

Figure

2.5

yields

a

binding

capacity

of

540

nmol

of

phosphoangiotensin/membrane (22 mm-diameter). Based on the 55 mol of TiO2 in the
membrane, the binding capacity is 126 mol phosphopeptides/gram of TiO2. This result agrees
reasonably well with a previous study that reported a binding capacity of 520 mol phenyl
phosphate/gram of 40 nm TiO2.[30] Phenyl phosphate is much smaller than phosphoangiotensin
and may give higher biding capacities
In most analytical applications, membrane loading is much less than saturation to avoid
the loss of desired analytes in the permeate. Defining dynamic capacity as the amount of binding
that occurs before the permeate concentration is 10 % of that in the feed; the dynamic
phosphoangiotensin-binding capacity is 365 nmol phosphopeptides per membrane. Moreover,
the dynamic capacity is the same at flow rates of 2 and 4 mL/min, suggesting kinetics and
diffusion do not limit binding prior to approaching saturation

72

Permeate Concentration (M)

25
20
15
10
5

0
0

5

10

15 20 25 30 35
Permeate Volume (mL)

40

45

50

Figure 2.5. Breakthrough curve for the passage of 20 M phosphoangiotensin (in loading buffer
I, 0.1% TFA) through a 22 mm-diameter nylon membrane modified with a (PSS/TiO2)2 film.
The flow rate through the membrane was 2 mL/min. Angiotensin served as an internal standard,
and the large error bars (standard deviation of 40%), result from the uncertainty inherent in
quantitative analysis based on MALDI-MS.

2.3.3 Phosphopeptide elution
In experiments that examined phosphopeptide elution and membrane reusability, 5 mL of
4 M phosphoangiotensin in loading buffer I was passed through the membrane (2 mL/min)
followed by 5 mL of washing buffer I and 5 mL of elution buffer at the same flow rate. Finally
the membrane was reconditioned with 10 mL loading buffer I, and the same experiment was
repeated nine times. The concentrations of phosphoangiotensin in the loading, rinsing and eluate
73

solutions were determined with MALDI-MS using angiotensin as an internal standard. (A
calibration curve accounted for small differences in the ionization efficiencies of angiotensin and
phosphoangiotensin).

Comparison of the amount of phosphoangiotensin removed from the

loading solution with the amount of phosphoangiotensin in the eluate gave an efficiency of
phosphoangiotensin elution between 80 and 100%. Unfortunately, the limited accuracy of the
MALDI-MS analyses does not allow us to know whether there is a small (10 to 20 %)
phosphopeptide loss during elution. To examine the stability of membrane performance, we
determined the elution efficiency over 10 experiments with the same membrane. Figure 2.6
shows that the fraction of phosphoangiotensin recovered in the eluate is stable at ~90%, which
suggests that there is no significant loss of TiO2 or buildup of phosphoangiotensin in the

Percent recovery

membrane during the experiment.
120
100
80
60
40
20
0

1

2

3

4

5

6

7

8

9

10

Trial number
Figure 2.6. Fraction of the phosphoangiotensin recovered from a loading solution after binding
and elution from a nylon membrane modified with a (PSS/TiO2)2 film. The same membrane was
employed repeatedly.
74

2.3.4 Phosphopeptide recovery
Although phosphoangiotensin recovery from single-peptide solutions (amount in
eluate/amount in loading solution) is around 90 %, recoveries from mixtures of digested proteins
may be much less because nonspecific binding may block adsorption sites. To quantitatively
examine recoveries in a more complex mixture, we employed H5 (CH3CH2COLFTGHPEpSLEK) and D5 (CD3CD2CO-LFTGHPEpSLEK) peptides, where the D5 peptide
spotted on the MALDI plate with the sample served as an internal standard (see Figure 2.7 for
the calibration curve).

We added 10 pmol of the target H5 peptide (CH3CH2CO-

LFTGHPEpSLEK) to a 150 pmol BSA digest in 100 L of loading buffer I, passed this mixture
through a nylon membrane modified with (PSS/TiO2)2 (Figure 2.8 shows mass spectra of the
solution before and after passing through the membrane), rinsed the membrane to remove nonspecifically bound peptides, and eluted the phosphopeptides with 30 L of elution buffer.
Comparison of the H5/D5 peptide signal intensity ratios for reference and eluate solutions
(Figure 2.9) indicates an overall H5 peptide recovery >70% in the eluate. The high recovery
occurs despite a 15-fold excess of BSA.

75

H5/D5 signal ratio

1.2

2
R²= 0.9974
R =0.9974

1
0.8
0.6
0.4
0.2
0
0

200

400

600

800

1000

Amount of H5 / fmol

Figure 2.7. Calibration curve showing the ratio of H5 to D5 peptide signals from MALDI-MS
analysis. The amount of D5 peptide spotted in all analyses was 1000 fmol, and the amount of H5
peptide varied.

76

a

b

a

H5:1393.45

b

H5:1393.45

600

800

1000

1200

1400 1600
m/z

1800

2000

2200

2400

Figure 2.8. MALDI mass spectra of 1 L of a solution containing 100 fmol H5 peptide and 1.5
2

pmol digested BSA (a) before and (b) after passing through a small membrane (0.02 cm )
modified with (PSS/TiO2)2.

77

D5:1398.45

a
H5:1393.45

b

D5:1398.45

H5:1393.45

1390

1393

1396

m/z

1399

1402

1405

Figure 2.9. MALDI mass spectra of (a) 500 fmol of D5 peptide internal standard with 333 fmol
of H5 peptide, and (b) 500 fmol of D5 internal standard plus 1 L of the eluate from enrichment
of 10 pmol H5 peptide into 30 L of elution buffer. For spectrum (b), 100% recovery would
correspond to spotting of 333 fmol, which should yield a spectrum similar to (a).

78

2.3.5 Phosphopeptide enrichment from a mixture of -casein, -casein, and
ovalbumin digests
In a test of enrichment from digests on only phosphoproteins, fifteen pmol of three
digested phosphoproteins were combined in 100 L of loading buffer I. The conventional
MALDI mass spectrum of 1 L (150 fmol of each protein) of this solution has weak
phosphopeptide signals compared to signals of non-phosphorylated peptides (Figure 2.10a).
Passage of the same solution through a TiO2-modified membrane both isolates and concentrates
phosphopeptides because binding occurs from 100 L of solution and elution requires only 10
L of eluent. The mass spectrum of the enriched sample (Figure 2.10b) contains 15 signals due
to phosphopeptides and in the few cases where the phosphopeptide signals are also present in the
loading-solution MALDI mass spectrum, signal intensities are an order of magnitude stronger
after enrichment. (The large number of peptides results from some missed and nonspecific
cleavages and variable alkylation.) Moreover, the spectrum of the enriched sample has only a
few weak signals (S/N <8) that stem from non-phosphorylated peptides.

79

aa



b b

o





o






1000

1500





2000



2500
m/z



3000

3500

4000

Figure 2.10. (a) MALDI mass spectrum of 1 L of loading buffer I containing 150 fmol (each)
of digested -casein, -casein and ovalbumin. This solution also contained 13 mM urea. (b)
Mass spectrum of the peptides collected from 100 L of the above solution by adsorption on a
nylon membrane modified with (PSS/TiO2)2, rinsing of the membrane with 100 L washing
buffer I, and elution of the peptides with 10 L of elution buffer. One L of the eluate was
spotted on the MALDI plate prior to addition of matrix.

80

2.3.6 Phosphopeptide enrichment from -casein, -casein, and ovalbumin
digests in a 100-fold excess of digested BSA
To mimic a modestly complex biological sample, we examined enrichment from 500 L
of loading buffer I containing 15 pmol (each) of digested -casein, -casein, and ovalbumin
along with 1500 pmol of digested BSA.

In this case, to reduce non-specific binding we

employed 2 % TFA in 1:1 water:ACN as the loading buffer.[33] The MALDI mass spectrum of
the loading solution (1 L of solution, 30 fmol of each phosphopeptide and 3 pmol of digested
BSA) reveals only one small phosphopeptide signal, along with many signals from BSA peptides
(Figure 2.11a). In contrast the mass spectrum of the enriched sample (Figure 2.11b) contains
signals from 19 phosphopeptides and no non-phosphorylated peptide signals with S/N >3. The
S/N for the phosphopeptides ranges from 3 to 50. Increases in the phosphopeptide S/N stem
from a 50-fold pre-concentration (assuming full recovery) and a decrease in ion suppression.
The S/N of the enriched sample is about half of that for enriched digests that do not contain BSA
(Figure 2.10), presumably because the higher TFA concentration decreases the efficiency of
phosphopeptide binding to the TiO2. (Table 2.1 contains m/z values, S/N, and sequences for the
peptides identified from Figure 2.11.)

81

a





b



  

  o
1000

1500





2000

o

2500
m/z





3000

3500

4000

Figure 2.11. (a) MALDI mass spectrum of 1 L of a loading solution containing 30 fmol (each)
of digested -casein, -casein and ovalbumin and 3 pmol of BSA in 260 mM urea. (b) Mass
spectrum of the phosphopeptides enriched from 500 L of the above loading solution by
adsorption on a (PSS/TiO2)2-modified nylon membrane, rinsing with 3 mL washing buffer II,
and recovery with 10 L of elution buffer. One L of the eluate was spotted on the MALDI
plate prior to addition of matrix.

82

Table 2.1. Phosphopeptides identified from the mass spectrum (Figure 2.11) of the membraneenriched peptides from tryptic digests of -casein, -casein, and ovalbumin in loading buffer II.
The loading solution contained a 100-fold excess of BSA. The m/z values, source proteins,
peptide sequences and S/N are listed.

m/z
1660.64
1703.64

protein
-casein
-casein

1717.64

-casein

1832.55
1879.45

-casein
-casein

1927.36
1952.64
1985.45

-casein
-casein
-casein

1995.73

-casein

2062.55
2089.55
2105.55

-casein
Ovalbumin
-casein

2119.55

-casein

2511.73
2966.73
3009.73
3024.82

ovalbumin
-casein
-casein
-casein

3122.82
3179.91

-casein
-casein

Peptide sequence
K.VPQLEIVPNpSAEER.L
K.VPQLEIVPNpSAEER.L
Carbamyl (N-term)
K.VPQLEIVPNpSAEER.L
Carbamidomethyl (E)
YLGEYLIVPNpSAEER
YLGEYLIVPNpSAEER
Carbamyl (N-term)
K.DIGpSEpSTEDQAMEDIK.Q
K.YKVPQLEIVPNpSAEER.L
K.DIGpSEpSTEDQAMEDIK.Q

Carbamidomethyl (D)
K.YKVPQLEIVPNpSAEER.L
Carbamyl (N-term)
K.FQpSEEQQQTEDELQDK.I
R.EVVGpSAEAGVDAASVSEEFR.A
K.FQpSEEQQQTEDELQDK.I
Carbamyl (N-term)
K.FQpSEEQQQTEDELQDK.I
Carbamidomethyl (E)
K.LPGFGDpSIEAQCGTSVNVHSSLR.D
R.ELEELNVPGEIVEpSLpSpSpSEESITR.I
R.NANEEEYSIGpSpSpSEEpSAEVATEEVK.I

A.RELEELNVPGEIVEpSLpSpSpSEESITR.I
(-H3PO4)
A.RELEELNVPGEIVEpSLpSpSpSEESITR.I
A.RELEELNVPGEIVEpSLpSpSpSEESITR.I
Carbamidomethyl (E)

83

S/N
23
4
6
3
15
11
13
7
11
50
5
14
10
9
24
6
5
19
3

2.4

Conclusions
Layer-by-layer modification of porous substrates with PSS/TiO2 nanoparticle films is a

convenient method for rapidly producing membranes that enrich phosphopeptides from smallvolume samples. The TiO2-modified membranes have a binding capacity of 540 nmol
phosphoangiotensin/membrane for a 22-mm diameter disc, and a capacity of 3 nmol in
membranes designed to handle small-volume (10’s of L) samples. Under sufficiently acidic
conditions (2% TFA), the membranes selectively enrich phosphopeptides from a digest
containing a 1:100 molar ratio of phosphoprotein to unphosphorylated protein.

Thus, the

membranes may prove useful in enriching phosphopeptides from modestly complex mixtures
such as fusion proteins or immunoprecipitates.

2.5

Acknowledgement

We are grateful to the U.S. National Institutes of Health (GM 080511 to M.B.) for funding this
work. I would like to thank Dr. Weihan Wang for help in the early stage and Dr. Amanda
Palumbo for providing the D5 and H5 peptides.

84

REFERRENCE

85

2.6

References

1.

Lee, M.; Goodbourn, S., Signalling from the Cell Surface to the Nucleus. In Regulation
of Gene Expression, Portland Press Ltd: London, 2001; Vol. 37, pp 71-85

2.

Cohen, P., The Regulation of Protein Function by Multisite Phosphorylation - a 25 Year
Update. Trends Biochem. Sci 2000, 25 (12), 596-601.

3.

Graves, J. D.; Krebs, E. G., Protein Phosphorylation and Signal Transduction. Pharmacol.
Ther. 1999, 82 (2-3), 111-121.

4.

Hunter, T., Signaling - 2000 and Beyond. Cell 2000, 100 (1), 113-127.

5.

Aebersold, R.; Mann, M., Mass Spectrometry-Based Proteomics. Nature 2003, 422
(6928), 198-207.

6.

Olsen, J. V.; Blagoev, B.; Gnad, F.; Macek, B.; Kumar, C.; Mortensen, P.; Mann, M.,
Global, in vivo, and Site-Specific Phosphorylation Dynamics in Signaling Networks. Cell
2006, 127 (3), 635-648.

7.

Thingholm, T. E.; Jensen, O. N.; Larsen, M. R., Analytical Strategies for
Phosphoproteomics. Proteomics 2009, 9 (6), 1451-1468.

8.

Witze, E. S.; Old, W. M.; Resing, K. A.; Ahn, N. G., Mapping Protein Post-Translational
Modifications with Mass Spectrometry. Nat. Methods 2007, 4 (10), 798-806.

9.

Pinkse, M. W. H.; Uitto, P. M.; Hilhorst, M. J.; Ooms, B.; Heck, A. J. R., Selective
Isolation at the Femtomole Level of Phosphopeptides from Proteolytic Digests Using 2dNanoLC-ESI-MS/MS and Titanium Oxide Precolumns. Anal. Chem. 2004, 76 (14),
3935-3943.

10.

Cantin, G. T.; Shock, T. R.; Park, S. K.; Madhani, H. D.; Yates, J. R., Optimizing TiO2Based Phosphopeptide Enrichment for Automated Multidimensional Liquid
Chromatography Coupled to Tandem Mass Spectrometry. Anal. Chem. 2007, 79 (12),
4666-4673.

86

11.

Thingholm, T. E.; Larsen, M. R.; Ingrell, C. R.; Kassem, M.; Jensen, O. N., TiO2-Based
Phosphoproteomic Analysis of the Plasma Membrane and the Effects of Phosphatase
Inhibitor Treatment. J. Proteome Res. 2008, 7 (8), 3304-3313.

12.

Pinkse, M. W. H.; Mohammed, S.; Gouw, L. W.; van Breukelen, B.; Vos, H. R.; Heck, A.
J. R., Highly Robust, Automated, and Sensitive on Line TiO2-Based Phosphoproteomics
Applied to Study Endogenous Phosphorylation in Drosophila Melanogaster. J. Proteome
Res. 2008, 7 (2), 687-697.

13.

Li, Y.; Xu, X. Q.; Qi, D. W.; Deng, C. H.; Yang, P. Y.; Zhang, X. M., Novel
Fe3O4@TiO2 Core-Shell Microspheres for Selective Enrichment of Phosphopeptides in
Phosphoproteome Analysis. J. Proteome Res. 2008, 7 (6), 2526-2538.

14.

Wang, H.; Duan, J.; Cheng, Q., Photocatalytically Patterned TiO2 Arrays for on-Plate
Selective Enrichment of Phosphopeptides and Direct MALDI MS Analysis. Anal. Chem.
2011, 83 (5), 1624-1631.

15.

Paradela, A.; Albar, J. P., Advances in the Analysis of Protein Phosphorylation. J.
Proteome Res. 2008, 7 (5), 1809-1818.

16.

Reinders, J.; Sickmann, A., State-of-the-Art in Phosphoproteomics. Proteomics 2005, 5
(16), 4052-4061.

17.

Leitner, A., Phosphopeptide Enrichment Using Metal Oxide Affinity Chromatography.
Trac-Trend Anal. Chem. 2010, 29 (2), 177-185.

18.

Han, G. H.; Ye, M. L.; Zou, H. F., Development of Phosphopeptide Enrichment
Techniques for Phosphoproteome Analysis. Analyst 2008, 133 (9), 1128-1138.

19.

Tichy, A.; Salovska, B.; Rehulka, P.; Klimentova, J.; Vavrova, J.; Stulik, J.; Hernychova,
L., Phosphoproteomics: Searching for a Needle in a Haystack. J. Proteome 2011, 74 (12),
2786-2797.

20.

Kweon, H. K.; Hå
kansson, K., Metal Oxide-Based Enrichment Combined with GasPhase Ion-Electron Reactions for Improved Mass Spectrometric Characterization of
Protein Phosphorylation. J. Proteome Res. 2008, 7 (2), 749-755.

87

21.

Carrascal, M.; Ovefletro, D.; Casas, V.; Gay, M.; Ablan, J., Phosphorylation Analysis of
Primary Human T Lymphocytes Using Sequential IMAC and Titanium Oxide
Enrichment. J. Proteome Res. 2008, 7 (12), 5167-5176.

22.

Dunn, D. J.; Reid, E. G.; Bruening, L. M., Techniques for Phosphopeptide Enrichment
Prior to Analysis by Mass Spectrometry. Mass Spectrom. Rev. 2010, 29 (1), 29-54.

23.

Connor, P. A.; McQuillan, A. J., Phosphate Adsorption onto TiO2 from Aqueous
Solutions: An in Situ Internal Reflection Infrared Spectroscopic Study. Langmuir 1999,
15 (8), 2916-2921.

24.

Larsen, M. R.; Thingholm, T. E.; Jensen, O. N.; Roepstorff, P.; Jorgensen, T. J. D.,
Highly Selective Enrichment of Phosphorylated Peptides from Peptide Mixtures Using
Titanium Dioxide Microcolumns. Mol. Cell. Proteomics 2005, 4 (7), 873-886.

25.

Tsai, C. F.; Wang, Y. T.; Chen, Y. R.; Lai, C. Y.; Lin, P. Y.; Pan, K. T.; Chen, J. Y.;
Khoo, K. H.; Chen, Y. J., Immobilized Metal Affinity Chromatography Revisited:
pH/Acid Control toward High Selectivity in Phosphoproteomics. J. Proteome Res. 2008,
7 (9), 4058-4069.

26.

Dotzauer, D. M.; Dai, J. H.; Sun, L.; Bruening, M. L., Catalytic Membranes Prepared
Using Layer-by-Layer Adsorption of Polyelectrolyte/Metal Nanoparticle Films in Porous
Supports. Nano Lett. 2006, 6 (10), 2268-2272.

27.

Dunn, J. D.; Igrisan, E. A.; Palumbo, A. M.; Reid, G. E.; Bruening, M. L.,
Phosphopeptide Enrichment Using MALDI Plates Modified with High-Capacity Polymer
Brushes. Anal. Chem. 2008, 80 (15), 5727-5735.

28.

Choi, W. Y.; Termin, A.; Hoffmann, M. R., The Role of Metal-Ion Dopants in QuantumSized TiO2 - Correlation between Photoreactivity and Charge-Carrier Recombination
Dynamics. J. Phys. Chem. 1994, 98 (51), 13669-13679.

29.

Lin, B.; Li, T.; Zhao, Y.; Huang, F. K.; Guo, L.; Feng, Y. Q., Preparation of a TiO2
Nanoparticle-Deposited Capillary Column by Liquid Phase Deposition and Its
Application in Phosphopeptide Analysis. J. Chromatogr. 2008, 1192 (1), 95-102.

30.

Liang, S. S.; Makamba, H.; Huang, S. Y.; Chen, S. H., Nano-Titanium Dioxide
Composites for the Enrichment of Phosphopeptides. J. Chromatogr. 2006, 1116 (1-2),
38-45.
88

31.

Hsieh, H. C.; Sheu, C.; Shi, F. K.; Li, D. T., Development of a Titanium Dioxide
Nanoparticle Pipette-Tip for the Selective Enrichment of Phosphorylated Peptides. J.
Chromatogr. 2007, 1165 (1-2), 128-135.

32.

Lewis, S. R.; Datta, S.; Gui, M.; Coker, E. L.; Huggins, F. E.; Daunert, S.; Bachas, L.;
Bhattacharyya, D., Reactive Nanostructured Membranes for Water Purification. Proc.
Natl. Acad. Sci. U. S. A. 2011, 108 (21), 8577-8582.

33.

Thingholm, T. E.; Jorgensen, T. J. D.; Jensen, O. N.; Larsen, M. R., Highly Selective
Enrichment of Phosphorylated Peptides Using Titanium Dioxide. Nat. Protoc. 2006, 1 (4),
1929-1935.

89

Chapter 3. Investigation of Phosphorylation in Tau Protein
and the Nuclear Factor Kappa B (NF-B) Pathway using
Phosphopeptide Enrichment withTiO2-modified Membranes
(Work on phosphorylation of Tau was submitted as part of an article in Analytical Chemistry)
Chapter 2 describes the fabrication of TiO2-modified nylon membranes and demonstrates
their enrichment performance with model phosphoprotein digests. This chapter presents initial
applications of these membranes for investigating phosphorylation of biologically important
proteins, including immunoprecipitated species and proteins phosphorylated in vitro and in vivo.
The goal is to demonstrate the strengths and limitations of the membranes in phosphopeptide
enrichment with biologically relevant mixtures.

3.1

Introduction
Phosphopeptide enrichment is a prerequisite step for bottom up studies of protein

phosphorylation because the sample complexity and low abundance of the target
phosphopeptides impede their identification with MS analysis.[1-5] As described in the previous
chapter, TiO2-modifed nylon membranes (22-mm diameter disk) have a binding capacity of 540
nmol of phosphopeptides and enrich phosphopeptides from a 100-fold excess of nonphosphorylated, digested protein. These properties should enable TiO2-modified membranes to
effectively enrich phosphopeptides from modestly complex biological mixtures such as digests
of fusion proteins, multi-protein complexes, or immunoprecipitates.

90

In contrast to global phosphoproteomics strategies that use SCX fractionation,
MOAC/IMAC enrichment, and HPLC-MS,[6-7] our interests focus on investigating
phosphorylation in proteins associated with a specific pathway or pathology, for instance the role
of tau protein phosphorylation in Alzheimer’s disease.[8-9] To demonstrate the utility of TiO2modified membranes, we examine phosphopeptide enrichment from a digest of tau protein that is
phosphorylated by glycogen synthase kinase-3GSK-3. The enrichment and analysis requires
only 8 g of protein, and the membrane eluent is amenable to analysis by both MALDI-MS and
ESI-MS to identify the number and location of phosphorylation sites. The combination of
membrane enrichment with tandem mass spectrometry reveals seven phosphorylation sites from
in vivo phosphorylated tau (p-tau) protein, which is associated with Alzheimer’s disease.
The NF-B pathway employs a series of protein complexes to alter DNA transcription in
almost all animal cells (Figure 3.1).[10] The signal transduction of NF-B facilitates the cellular
response to various stimuli, such as cytokines, environmental stress, UV-irradiation, radicals,
antigens and bacterial infection.[11-12] Malfunction of the NF-B pathway may lead to
inflammation or immune disease, and even cancer.[13-14] Studies of phosphorylation related to
the key transcriptional factor p65 (RelA) under the stimulation of tumor necrosis factor (TNF-)
will provide insight to further understand the NF-B pathway.[15] In this chapter, the
combination of membrane enrichment with MALDI MS/MS reveals eight phosphopeptides from
p65-associated proteins immunoprecipitated from human acute monocytic leukemia (THP-1) cell
nuclear extracts.
The inhibitor of kappa-B-alpha (IB) is also an important protein in the NF-B pathway,
as IB conserves and inhibits transcriptional factors p65 and p50 in the cytoplasm.[16] In
addition to the well-known Inhibitor of kappa-B kinase beta (IKK), some studies suggest that
91

the G-protein coupled receptor kinase-5 (GRK5) also mediates the activation of the NF-B
pathway.[17-18] Even without enrichment, our LC-MS/MS analysis reveals two phosphorylation
sites (Ser32 and 36) from a digest of IB that is in vitro phosphorylated by GRK5.
Nuclear Factor Kappa B (NF-kB) pathway
Signal (tumor necrosis factor : TNF-)
Receptor

Cell membrane

IB

P P

IKK

Cytoplasm

IB kinase

Inhibited
p65

Cell survivor
Inflammation
Immunity

P P

Proteasome
degradation

p50

Nucleus
Gene
transcription

Transcriptional factor

translocation
Nuclear
DNA

mRNA

Figure 3.1. Cartoon of the NF-B pathway under the stimulation of TNF-After the receptor
outside the cellular membrane receives a signal (such as the cytokine TNF-the released IKK
will phosphorylate the inhibited IB-p65/p50 complex in the cytoplasm. The phosphorylated
IB-p65/p50 complex tends to undergo proteasome degradation and release the two
transcriptional factors p65 and p50. The free transcriptional factors translocate into the nucleus
and bind to the nuclear DNA to create different transcriptional mRNA. The normal NF-B
pathway would provide the right response for cell survival/apoptosis and immune response to
outside stimulation.
92

3.2

Experimental

3.2.1 Materials

and

solutions,

membranes

and

holders,

membrane

modification and protein digestion
See chapter 2 for details in this area.

3.2.2 Purification of p-tau
Protein expression and purification were performed by Dr. Dexin Sui from Prof. MinHao Kuo’s group. Tau protein was overexpressed in E. Coli. A single plasmid contained genes
for His6FosGSK3 and Juntau fusion proteins. Bacterial cells in buffer (20 mM Tris, 100 mM
NaCl and 2 mM orthovanadate, pH 8, production of a 0.1 g bacterial pellet requires 8 mL buffer)
were sonicated, and the sample supernatant was saved after centrifuging at 13000 rpm for 30 min
at 4 º The supernatant was loaded on a Ni-NTA column and washed with buffer prior to
C.
elution with a 30 - 300 mM imidazole gradient. Eluate fractions were combined and concentrated
in a 30 kDa spin column to a volume of 6 - 10 mL. One mL of the concentrated sample was
loaded onto a size exclusion column (Superdex 200 10/300 GL) coupled to an fast protein liquid
chromatography (FPLC) system (Amersham pharmacy biotech AKTA P-920). The eluent was
20 mM Tris and 100 mM NaCl at 0.4 mL/min. Fractions were collected every 1 mL and
monitored by their absorbance at 280 nm. UV-absorbing fractions for a given peak were
combined and concentrated with a 30 kDa spin column at 4000 rpm at 4 º The final protein
C.
concentration was estimated from the UV absorbance at 280 nm, and these samples were
digested and employed in MS studies.

93

3.2.3 Enrichment of phosphopeptides from digested p-tau
After in-solution digestion of 100 g of purified protein using the protocol in Chapter 2
(page 65), 160 pmol of p-tau digest was dissolved in 200 L of loading buffer II (2 % TFA in 1:1
water:ACN) and passed through a small-scale TiO2-modified membrane at 10 L/min. The
membrane was washed with 200 L of washing buffer II (2 % TFA in 1:1 water:ACN) at the
same flow rate, and phosphopeptides were eluted with 50 L of elution buffer (1 % NH4OH in
1:1 water:ACN) at 5 L/min.

3.2.4 Immunoprecipitation of p65 from THP-1 cells
The THP-Cells (from ATCC, Manassas, VA) were cultured in RPMI medium 1640
(Invitrogen) at 37 º with 5% CO2 and ambient O2 until reaching a concentration of 9.6 x 105/mL
C
in flow cytometry. The cells were treated with TNF-at a final concentration of 10 ng/mLfor 30
min before harvest. The cell suspension was centrifuged (500g) for 5 min at 4 º and after
C,
rinsing with 10 mL ice-cold phosphate buffered saline (PBS), the suspension was centrifuged
again and the cells were lysed with 1.5 mL lysis buffer (10 mM HEPES pH=8.0, 1.5 mM MgCl2
and 10 mM KCl) for 30 min on ice. The pellet from subsequent centrifugation was further
extracted with 60 L nuclear extraction buffer (20 mM HEPES pH=8.0, 1.5 mM MgCl2, 0.42 M
NaCl, 0.2 mM EDTA and 25 % glycerol) on a shaker (2500 rpm) for 30 min at 4 º The final
C.
total protein concentration was ~6 g/L according to a Bradford assay. The phosphorylation of
p65 was confirmed by Western blotting (WB) using a primary p65 polyclonal antibody (C-20
from Santa Cruz Biotechnology). Twenty plates of cells were treated in parallel, and 1000 g of
total cell extract was pooled into 800 L PBS buffer and mixed with 30 L covalently cross94

linked p65 antibody agorose (C20 from Santa Cruz Biotechnology) in 200 L nuclear extraction
buffer. The IP efficiency was probed using WB with a p65 antibody (C22B4 from Cell Signaling
Technology, Boston, MA). Protein on the 30-L antibody beads was digested according to the
protocol for digesting 100 g BSA (chapter 2, page 65), starting with the addition of the beads to
the urea solution. This overall procedure was initially developed by Dr. Amanda Palumbo.[19]

3.2.5 Enrichment of on-bead digests of immunoprecipitated p65 and
associated proteins
Sixteen L of the p65 on-bead digest was dissolved in 200 L of loading buffer I (0.1 %
TFA in 1:1 water:ACN), and 5 pmol of ovalbumin digest was added to the solution. The sample
2

was passed through the small membrane (0.02 cm ) at a flow rate of 10 L/min, and the
membrane was rinsed with 200 L washing buffer I (0.1 % TFA in 1:1 water:ACN) at the same
flow rate.

Finally 10 L of (1 % H3PO4 in 1:1 water:ACN), was used to elute the

phosphopeptide from the membrane at a flow rate of 2 L/min.

3.2.6 In vitro phosphorylation of IB by GRK5 and IKK
The IB and protein kinases were kindly provided by Prof. Parameswaran. The 20 L
reaction system contained 2.6 L of 0.049 g/L IB, 0.5 L of 10 mM ATP, 0.25 L of 0.22
g/L IKKor 0.26 L of 0.17 g/L GRK5 (from Cell Signaling Technology, Boston, MA),
2.5 L kinase buffer (from Cell Signaling Technology, Boston, MA) and H2O (bring up to 20
L). The in vitro phosphorylation was carried for 30 min at room temperature and stopped by
addition of 5 L of 2% SDS buffer. The IB phosphorylation by GRK5 and IKKwas

95

confirmed by WB analysis. The system was scaled up to 5 g of IB for the LC-MS/MS
analysis. Five g of IB with and without phosphorylation by GRK5 was digested in solution
prior to the LC-MS/MS analysis.

3.2.7 Mass Spectrometry and Data Analysis
3.2.7.1

MS method for p-tau phosphorylation study
The MALDI analysis was the same as described in Chapter 2. For ESI-MS analyses, 40

µL samples in 1% acetic acid, 49% H2O and 50% methanol were loaded into a Whatman
multichem 96-well plate (Sigma Aldrich, St. Louis, MO) and sealed with Teflon Ultra Thin
Sealing Tape (Analytical Sales and Services, Prompton Plains, NJ). Then, using an Advion
Triversa Nanomate nano-electrospray ionization (nESI) source (Advion Ithaca, NY) with a spray
voltage of 1.4 kV and a gas pressure of 1.0 psi, samples were introduced into a high resolution
accurate mass LTQ Orbitrap Velos mass spectrometer (Thermo Fisher Scientific, San Jose, CA)
equipped with a dual pressure ion trap, Higher-Energy Collisional Dissociation (HCD) cell and
Electron Transfer Dissociation (ETD). The ion source interface settings (inlet temperature of 200
o

C and S-Lens value of 60 %) were optimized to maximize the sensitivity of the precursor ions

while minimizing ‘in-source’ fragmentation. High resolution mass spectra, as well as all ion trap
n

CID-, HCD- and ETD-MS/MS and -MS spectra, were acquired in positive ionization mode
using the FT analyzer operating at 100,000 resolving power. Spectra were typically signal
averaged for 30-100 scans. Peptide precursor ions were mass-selected using an isolation width of
5 Da. CID was performed using default activation parameters (q=0.25, activation time 10 msec)
with a normalized collision energy of 30. HCD was performed using default activation
parameters, with a collision energy of 20-40 and a low mass cutoff of 100. For ETD, the
96

automatic gain control value and the maximum injection time of the reagent ion (fluoranthene
5

anion) were set as 1.0 × 10 and 100 ms respectively, and the reaction time was optimized as 50
ms for +3 charge state. Assignment of peptides resulting from protein digestion was performed
manually by mass fingerprinting or by MS/MS. Phosphorylation sites were manually assigned by
matching MS/MS product ions with in silico peptide fragments generated using
ProteinProspector (http://prospector.ucsf.edu).

3.2.7.2

MS method for p65 phosphorylation study
To increase the amount of material spotted on the MALDI plate, 5 L of eluate (in 1 L

increments that were allowed to dry) was spotted onto the same well prior to addition of 1L of
2,5-DHB matrix. The additional spotting gives stronger signal intensity and more precursor ions
for MS/MS than the traditional 1L spotting method. Ion trap CID-MS/MS was performed on
mass-selected precursor ions, and MS/MS spectra shown were typically collected for about 1
hour to average 3000 scans. MS/MS spectra were transformed to DTA files by Bioworks 3.3
(Thermo Fisher Scientific), and searched using the Mascot algorithm (Matrix Science) under the
NCBInr data base with taxonomy as homo sapiens (human) for p65 IP proteins. The
identifications and determined phosphorylation sites were also confirmed manually.

3.2.7.3

MS method for IB phosphorylation study
These analyses were performed by Dr. Xiao Zhou from Prof. Gavin E. Reid’s group. Five

L of digested IB at a concentration of 0.2 pmol/L in 3% acetic acid/5% ACN was loaded
from a Paradigm AS1 autosampler (from Michrom Bioresources, CA) onto a peptide CapTrap
(from Michrom Bioresources, CA) at a flow rate of 15 L/min using loading buffer 0.1% TFA/ 2%

97

ACN. After 5 min of loading, the peptides concentrated on the CapTrap were eluted onto a 200
m id × 50 mm fused silica column packed with Magic C18AQ (3 m particlefrom Michrom
Bioresources, CA) at a flow rate of 2 L/min using a linear 45 min gradient from 95% solvent A
(0.1% FA in H2O) to 50% solvent B (0.1% FA in ACN). After gradient separation, the system
was cleaned with 80% solvent B for 5 min followed by equilibrium with 95% solvent A for 5
min.
The mass spectrometer was programmed to operate in a data-dependent acquisition mode.
In the full scan analysis, the five most intense ion signals (the signal threshold was set at 1.0 ×
4

10 counts) from the survey MS scan were isolated with an isolation window of 2.0 m/z. During
4

CID-MS/MS, when an AGC target value of 1.0 × 10 was reached within 100 ms maximum
injection time in the ion trap, the isolated ions were fragmented with 35 normalized collision
energy using a 10 ms activation time and an activation q value of 0.25.

3.3

Results and discussion

3.3.1 Enrichment of p-tau digests
The studies in the previous chapter suggest that TiO2-modified membranes are attractive
for enriching phosphopeptides from modestly complex samples such as digests of complexes
containing fusion proteins. To test the membranes with a phosphoprotein with more direct
relevance to biology, we enrich phosphopeptides from a digest of hyperphosphorylated tau (ptau). Abundant in neurons, tau is a very soluble microtubule-associated protein that along with
tubulin assembles and stabilizes the microtubule cytoskeleton.[20] In many neurodegenerative
conditions such as Alzheimer’s disease, hyperphosphorylated tau dissociates from the
98

microtubule and aggregates into paired helical filaments and neurofibrillary tangles,[8] whose
abundance correlates well with the severity of neurodegeneration of Alzheimer's disease patients.
Thus, compounds that block or reverse tau aggregation may have therapeutic value in treating
Alzheimer's disease. Development of such potential drugs will benefit from a better
understanding of the sites of p-tau phosphorylation.
One of the candidate kinases for tau hyperphosphorylation in Alzheimer’s disease is
GSK-3. In vitro, GSK-3catalyzes tau phosphorylation that facilitates the formation of tanglelike aggregates.[9, 21-23] To prepare the p-tau subjects for digestion and our TiO2 enrichment,
Prof. Kuo and Dr. Sui employed a protein interaction module-assisted catalysis technique in
which GSK-3 and tau are expressed as His6-FosB and JunD fusion proteins, respectively
(Figure 3.2). The formation of a stable FosB-JunD leucine zipper complex in E. coli leads to
high-efficiency phosphorylation of tau. Subsequently, affinity-trapping of the His-tag with a NiNTA column enables isolation of the complex, which in this study was further purified through
size-exclusion FPLC. Selected FPLC fractions containing GSK-3p-tau and a small number of
bacterial proteins were pooled for MS. It should be noted that the complexity of these samples is
similar to our model solution containing phosphoproteins in excess BSA.

His6-Fos-GSK
His6-Fos-GSK

Phosphorylation

Jun-Tev-Tau

Jun-Tev-Tau
Figure 3.2. Scheme of the protein interaction module-assisted phosphorylation. The leucine
zipper Fos and Jun bring the enzyme and substrate together after they are co-produced in cells,
and phosphorylation occurs in vivo.
99

Figure 3.3a shows the MALDI mass spectrum of a digested FPLC fraction that contained
p-tau. The peptide signals stem from GSK-3p-tau, and unknown proteins that co-purify with
the complex. The sample also contains 0.12 M urea and 10 mM NaCl, which decrease the S/N,
so the highest S/N is 20 for a peptide from GSK-3Overall, the spectrum reveals 63 peptides
whose signals are above 10 % of the highest signal, but none of these are phosphopeptides.
Using the TiO2-modified membranes with loading buffer II, we capture the
phosphopeptides from 200 L of the same digested FPLC fraction and then recover these
peptides in 50 L of elution buffer. The mass spectrum of 1 L of the eluate spotted on a
MALDI plate (Figure 3.3b) shows a much simpler pattern than in Figure 3.3a, and the S/N is as
high as 100 for a phosphopeptide from p-tau. The TiO2-modified nylon membrane isolates,
concentrates (4-fold) and desalts the phosphopeptides to enable their detection. We identify
seven phosphorylation sites from three phosphopeptides, P1, P3, and P4 (P2 results from insource loss of 98 Da from P3.). Even the strongest phosphopeptide signal, P3 at 2455.00 m/z, is
not present in the mass spectrum of the membrane-loading solution, presumably because of
suppression. The enriched mixture also shows signals from four unphosphorylated peptides
(labeled as n in spectrum b). These signals are also present in Figure 5a, but they do not stem
from either p-tau or GSK-3 A control experiment with tau digests without GSK3stimulation (Figure 3.4) shows no detectable phosphopeptides after enrichment.

100

a

b
P1:195pSGYpSSPGSPGTPGSR
P3:386TDHGAEIVYKpSPVVpSGDTpSPR
P4:407HLpSNVSSTGSIDMVDpSPQL
ATLADESASLAK
n: unphosphorylated peptide

P2

P1

1000

1500

2000

m/z

P3

nn
n n

2500

P4

3000

Figure 3.3. (a) MALDI mass spectrum of a digested FPLC fraction that contained approximately
800 fmol of GSK-3-p-tau complex along with associated proteins; (b) The corresponding mass
spectrum of peptides enriched from the above solution by adsorption on a TiO2-modified nylon
membrane and subsequent elution. The figure shows the sequences of 4 phosphopeptides, one of
which, P2, results from in-source P3 dephosphorylation. Four major unphosphorylated peptides
are labeled with n.
101

a

a

b

b

1000

1500

2000

2500
m/z

3000

3500

4000

Figure 3.4. (a) MALDI mass spectrum of an FPLC fraction containing approximately 800 fmol
of digested tau protein and associated proteins. (b) The corresponding mass spectrum of
phosphopeptides enriched from the above solution by adsorption on a TiO2-modified nylon
membrane and elution. The spectra reveal no phosphopeptides, confirming that tau protein is not
phosphorylated without GSK3 catalysis.

To identify the phosphorylation sites on p-tau, we employed ESI-Orbitrap-MS coupled
n

with CID-, HCD-, or ETD-MS/MS and/or MSn fragmentation. In CID-MS (Figure 3.5) the
dominant sequential loss of phosphate under consecutive CID fragmentations confirms that
386

TDHGAEIVYKpSPVVpSGDTpSPR has three phosphate groups. The CID MS/MS spectrum

gives 87 fragment ions resulting from breaking of 17 out of 20 peptide bonds. The HCD MS/MS
spectrum shows 93 fragment ions from the breakage of 18 out of 20 peptide bonds (Figure 3.6).
The higher energy causes more internal fragmentation, leading to additional small fragments at a
low mass range compared to CID analysis. Both techniques provide high confidence for peptide
102

identification. However, the potential for competing losses of 98 Da (either directly as H3PO4
from a phosphorylated Serine or Threonine residue, or as the combined losses of 80 Da (HPO3)
from a phosphorylated Serine or Threonine residue and 18 Da (H2O) from a non-phosphorylated
Serine or Threonine residue) from either the precursor ion or from the b- and y-type product ions
may result in ambiguity in assigning the sites of phosphorylation within the peptide sequence.[24]
In contrast, retention of phosphate groups during ETD [25-26] allows confident assignment of
phosphorylation sites (Figure 3.7). Two other phosphopeptides were identified using HCD
analysis and the phosphate groups were assigned as

195

pSGYpSSPGSPGTPGSR and

407

HLpSNVSSTGSIDMVDpSPQLATLADESASLAK (see mass spectra in Figure 3.8). Overall,

we identified 7 phosphorylation sites (S195, S198, S396, S400, S404 S409 and S422) using MS
analysis following enrichment with the TiO2-modified nylon membrane.

103

386T

b172+

b112+

D H G A E I V Y K pS P V V pS G D T pS P R
y7

y11

a

y3

[M+3H] ∆3+
3+
[M+3H] Ƽ

[M+3H] ∆∆3+
3+
[M+3H] ∆∆º

y2

y3

b8

b112+

b

b17∆2+ b b19∆∆2+
9
y18∆2+ ∆º
y7 b172+
b19 2+ y
y8
b10 10 y11∆

y11

[M+3H] ∆∆3+
3+
[M+3H] ∆∆º
y11∆∆2+b

y2

y11∆∆∆2+
y4∆ y10∆2+

c

y11

∆∆2+

y11∆∆∆2+

y2
400

11

Ƽ
2+

b11∆2+
y11∆2+

b8

b ∆∆2+
b17∆2+ 19
y18∆2+
∆
b9
y8
y ∆
10

b10
y11∆∆

[M+3H] ∆∆∆3+
2+
b11Ƽ
∆2+
b11

b10

3+
[M+3H] ∆∆∆º
y19∆∆3+

600

b19∆∆∆2+ y10∆
y18∆∆2+
b9

800
m/z

y11∆∆

1000

Figure 3.5. ESI-Orbitrap multistage CID-MSn of the [M+3H]
386

1200
3+

1400

precursor ion of
3

TDHGAEIVYKpSPVVpSGDTpSPR (m/z 819.34). (a) CID-MS/MS, (b) CID-MS of the -98
4

Da product ion from panel A (m/z 786.48), (c) CID-MS of the -98 Da product ion (m/z 753.98)
from panel B. The sequences show cleavage sites for formation of b and y ions. （∆=-98 Da, o=18 Da）
104

y2
386T

D H G A E I V Y K pS P V V pS G D T pS P R
y7

y11

y3

[M+3H]∆3+
3+
[M+3H]Ƽ

[M+3H]3+

[M+3H]∆∆3+
K

b3
y1 b2

y11∆∆2+
∆2+
y3 b5 b11 y11∆2+
y4∆ y4 b6 y6 b7
b
4

b8

b17∆2+ b ∆2+ b
19
10
y7 y8 b9 y10∆ y10 y11∆ y11

100 200 300 400 500 600 700 800 900 1000 1100 1200 1300
m/z
Figure 3.6. ESI-Orbitrap HCD-MS/MS spectra of the [M+3H]
386

3+

precursor ion of

TDHGAEIVYKpSPVVpSGDTpSPR (m/z 819.34). The sequences show cleavage sites for

formation of b and y ions. （∆=-98 Da, o=-18 Da）

105

T D H G A E I V Y K pS P V V pS G D T pS P R
Z+13

[M+3H]3+

[M+3H]2+∙
C4
Z+13

C5

C-110
C8 Z+18
Z+15
Z19
C6
Z+16
Z+17 C9
C-17
Z+1
9

400 500 600 700 800 900 1000 1100 1200

m/z

Figure 3.7. ESI-Orbitrap ETD-MS/MS spectrum of the [M+3H]
386

3+

precursor ion of

TDHGAEIVYKpSPVVpSGDTpSPR (m/z 819.34). The sequence shows cleavage sites for

formation of c and z ions.

106

b
195pS G 3 pS S
Y

a
Y

PGSPGTPGSR
Experimental Mass: 1553.564
Theoretical Mass: 1553.567

y7
a3∆ ∆ y4
b3

y4*

y1
b2∆

b3

b5

y10

∆∆º

y12Ƽ
y6 y7* y y9
8 y ºy11º
b5 y y7º
∆
10
y8º
5
y11 y12
∆∆

200

400
600
800
1000
1200
1400
1600
m/z
b5
407H L pS N V S S T G S I D M V D pS P Q L A T L A D E S A S L A K
y16
b PQ
Experimental Mass: 3403.530
y10
Theoretical Mass:

3403.538

y12
y13

y9

y6 b5
b2 PQL
y2
y3

y1

200
Figure

3.8.

y4
y5

400

ESI-Orbitrap

b5∆

600

y11

y8
y162+
y7 b6º
b6
800

HCD-MS/MS

m/z
of

195

y14

1000
(a)

the

1200
[M+2H]

1400
2+

pSGYpSSPGSPGTPGSR (m/z 778.00) and (b) the [M+3H]
107

b14º y16

3+

1600

precursor

ion

of

precursor ion of

407

HLpSNVSSTGSIDMVDpSPQLATLADESASLAK (m/z 1136.00).

The figure lists the

experimental and theoretical masses of the peptides, and the assigned sequences and
phosphorylation sites. （∆=-98 Da, o=-18 Da and *=-17 Da）
Multiple approaches such as MS, WB, and Edman degradation have previously revealed
more than 40 phosphorylation sites in different tau isoforms,[27] making it particularly
challenging to map all phosphorylation sites of a given p-tau species. However, as Figure 3.3b
shows, the signal from the

386

TDHGAEIVYKpSPVVpSGDTpSPR (S396, S400 and S404)

peptide dominates the spectrum of the enriched peptides. There are three possible reasons for this
dominance. First, the peptide has a particularly high ionization efficiency. Given the presence of
3 phosphates, this seems unlikely for analysis in the positive ion mode. Second, the membrane
selectively enriches the tri-phosphorylated peptide. However, TiO2 usually shows a bias toward
mono-phosphorylated peptides.[19, 28] Third, and the most likely of the three, the stoichiometry
of

386

TDHGAEIVYKpSPVVpSGDTpSPR phosphorylation may surpass all other tryptic

phosphopeptides of p-tau. Using traditional methods of preparing p-tau with a commercial GSK3, the Gamblin group phosphorylated tau with 3 moles of phosphate incorporated per mole of
tau. Similarly, the Reynolds group reports 2-4 moles phosphate/mole of Tau.[22, 29] They also
find a S396-S400-S404 tri-phosphorylated peptide. We thus suspect that these three residues are
the most preferred targets for GSK-3. This phenomenon is also consistent with the observation
that GSK-3 activity can be enhanced by pre-existing, priming phosphorylation that lies 4
residues C-terminal to the next phosphorylation site.[30] On the other hand, whether any of these
three phosphorylation sites plays a key role in Alzheimer’s disease remains to be assessed.
Further refinement of the phosphorylation, sample preparation, and enrichment procedures will
likely yield more comprehensive mapping of GSK-3-phosphorylated residues in p-tau.
108

Identification of the most dominant and other sub-stoichiometric phosphorylation sites may
prove useful in designing drugs to prevent the pathological hyperphosphorylation of tau and even
the appearance of the neurofibrillary tangles.[28]

3.3.2 Phosphopeptide

enrichment

from

on-bead

digests

of

immunoprecipitated p65 and associated proteins
Immunoprecipitated proteins are somewhat similar in complexity to the fusion proteins
described above. In addition to peptides from the specific antigen protein, tryptic digests of
immunoprecipitates contain peptides from associated proteins, the antibody, and nonspecifically
adsorbed proteins. We examined enrichment of immunoprecipitated p65 and its associated
proteins. The heterodimeric p65-p50 complex is the most abundant member of the NF-family
of transcription factors present in almost all animal cell types.[31] These factors help regulate
many biological processes such as inflammation, immunity and tumor genesis,[32-35] and the
NF-pathway is mainly controlled by PTMs, especially phosphorylation.[36-37] Thus, we
hope to identify phosphorylated p65 or p65-associated proteins in the nuclear extract of cells
treated with a stimulating agent, TNF-.

109

Tumor necrosis factor (TNF-) treatment
Lysis
Human acute monocytic leukemia
(THP-1) cells

p65 C-20 antibody
agarose conjugate
Immunoprecipitation

1 mg nuclear
extracts

MS
analysis

On bead
digestion

Membrane
enrichment
Phosphorylation sites

m/z

Scheme 3.1 Workflow of the p65 phosphorylation study. The THP-1 cells were treated by TNFand 1 mg (based on a Bradford assay) of nuclear-extract proteins obtained from cell lysis was
immunoprecipitated by a p65 C-20 antibody-agarose conjugate. After on bead digestion, the
phosphopeptides were enriched by a TiO2-modifed membrane, followed by MS-analysis.
The p65 and associated proteins were initially immunoprecipitated from a nuclear extract
of TNF- stimulated THP-1 cells (Scheme 3.1). Although WB clearly shows the presence of
p65 in the treated cells and confirms the efficiency of IP, (Figure 3.9) we do not know how much
p65 or associated phosphoprotein is present in the immunoprecipitated protein. Tryptic digestion
was performed with the proteins still attached to the affinity beads, and 16 L of digest was dried
and dissolved in 200 L of loading buffer I. To assess whether the membrane enriches
phosphopeptides from this specific sample, we also add 5 pmol digested ovalbumin into the 200L solution prior to the enrichment procedure, which includes passing the digest through the
membrane, rinsing, and elution with 10 L of 1% H3PO4 in 1:1 water:ACN. We observed that
when the sample amount is low, the NH4OH elution leads to matrix cluster ions at a low mass

110

range in MALDI-MS. To avoid these cluster ions, we switched to an H3PO4 eluent that provides
high elution efficient and alleviates the cluster problem.

Marker

A

75 kD

B
Figure 3.9. The WB analysis of p65 using
primary polyclonal p65 antibody (C22B4).

50 kD

Lane A is the 25 g nuclear extracts before IP,
and lane B is the 25 nuclear extracts after IP.
The red circle highlights the p65.

37 kD
The MALDI mass spectrum of the digested and enriched IP contains an ovalbumin
phosphopeptide signal at m/z 2089 (Figure 3.10), and 8 other phosphopeptide signals due to the
p65 digest. A neutral loss of 98 Da in CID MS/MS spectra serves as the diagnostic marker for
phosphopeptides.[38]

111

*
**

100%=4.01E3
P65 digests and 2.5 pmol Ovalbumin

**
*
*
*

*

1000

1500

2000

2500
m/z

3000

3500

4000

Figure 3.10. MALDI mass spectrum of a membrane-enriched digest of immunoprecipitated p65
and associated proteins. Sixteen L of p65 on-bead dried digest and 5 pmol ovalbumin digest
were dissolved in 200 L loading buffer I, then passed through the TiO2-modified membrane,
and phosphopeptides were eluted in 10 L of 1% H3PO4 in 1:1 water:ACN. Five L of eluent
was spotted on the same sample well prior to MALDI-MS. The red stars represent the
phosphopeptides and the blue star is the phosphopeptide from the ovalbumin digest.
Interestingly, none of the phosphopeptides come from p65 itself. This most likely occurs
because digestion of p65 does not produce phosphopeptides in the mass range we employ (m/z
1000-4000). Using the ProteinProspector MS-Digest Program (with no missed cleavages), we
generate the potential p65 tryptic peptides that contain known phosphorylation sites.
Unfortunately, only one of these peptides has a mass between 800-4000 Da, and we cannot be

112

sure that this particular peptide is phosphorylated in our samples.[39] In retrospect, p65 is not the
best choice for verifying this technique.
The immunoprecipitates do contain eight phosphopeptides from the P1 or P2 isoform of
the 60S acidic ribosomal protein (Ribosomal), which is likely associated with p65. Table 3.1
shows the m/z values and corresponding sequences of each peptide. The peptide at m/z 2029 is
the mono-phosphopeptide (KEESEEpSDDDMGFGLFD), while the peptide at m/z 2109 is the
diphosphopeptide with same sequence (KEEpSEEpSDDDMGFGLFD). Figures 3.11a and 3.11b
show the MS/MS spectra of these peptides. The other phosphopeptides listed in Table 3.1 are
peptides sharing the same sequence (KEESEESDDDMGFGLFD) but with modifications on
methionine (Met) during alkylation.[40-42] The peptide at m/z 1734 is truncated from the
peptide at m/z 2166, which may result from non-specific cleavage by trypsin, chymotrypsin
proteolysis or proteolytic processing during sample preparation. We are unable to obtain a
sequence for the peptide at m/z 1947, but the CID MS/MS spectrum gives a predominant neutral
loss of 98 Da, which is an indication of a phosphopeptide ion. Additionally, at the low mass
range, the CID MS/MS of this peptide contains some product ions identical to those resulting
from the peptides with m/z 2029 and 1981. Thus, the peptide with m/z 1947 may share some of
the sequence of the other peptides and contain a modification that we did not discover.

113

a

b8
y5

b
Ƽ 8

b 9∆ b 9

800

1000

b10

∆

b10
b11∆ b11

1200

y14∆
b13∆ b13

1400
m/z

b16∆ ∆
y16
b15∆
b15 y16

1600

1800

y12
b6∆

800

b8

∆

∆

b9∆ b10∆ b11∆
b8º b
b
b10
y12 11
9

1000

1200

1400
m/z

b15∆
b14º
∆∆
y14
b16∆ b16
∆
∆
b13
y15
y16º
b13
b15

1600

1800

Figure 3.11. MALDI CID-MS/MS product-ion spectra of (a) the [M+H]
KEESEEpSDDDMGFGLFD (m/z 2029) and

(b) the [M+H]

º
2000

∆

b6
K E E pS E E pS D D D M G F G L F D
y6

b

600

∆
b16º

b10º

b6

600

y6

∆

b6
K E E S E E pS D D D M G F G L F D
y5

+

2000
+

º

2200

precursor ion of

precursor ion of

KEEpSEEpSDDDMGFGLFD (m/z 2109). The sequences show cleavage sites for formation of b
and y ions. （∆=-98 Da, o=-18 Da）

114

Table 3.1. Phosphopeptides identified from membrane-enriched tryptic digests of the nuclear
extracts from TNF--treated THP-1 cells. The m/z value, peptide sequence and source are listed.

m/z
+
[M+H]
1734.8
1947.5
1981.6
2029.6
2061.6
2109.5
2125.6
2166.8

Peptide sequence
KEEpSEEpSDDDMGF
(1677.5+57: S-carbamidomethylmethionine)
Not sequenced
KEESEEpSDDDMGFGLFD
(2029-48: loss of 2-(methylthio)acetamide )
KEESEEpSDDDMGFGLFD
KEEpSEEpSDDDMGFGLFD
(2109-48: loss of 2-(methylthio)acetamide)
KEEpSEEpSDDDMGFGLFD
KEEpSEEpSDDDMGFGLFD
(2109.5+16:Oxidation of methionine)
KEEpSEEpSDDDMGFGLFD
(2109.5+57: S-carbamidomethylmethionine)

Source
Ribosomal
Ribosomal
Ribosomal
Ribosomal
Ribosomal
Ribosomal
Ribosomal

These results suggest that the ribosomal protein associates with p65, although it is
possible that it non-specifically interacts with the antibody beads. Ribosomal protein is involved
in translational processes, and a study by Bouwmeester and coworkers indicates that nine
ribosomal proteins and four other proteins are involved in ribosomal function with p65.[43-44]
Thus it is highly likely that the Ribosomal protein associates with p65 during the IP process.

3.3.3 Analysis of in vitro phosphorylation of IB by GRK5
Recent studies by the Parameswaran group at Michigan State University suggest that
similar to IKK, GRK5 can also mediate the activation of the NF-pathway via
phosphorylation of IB.[17-18] The mutation of Ser 32 and 36, which are sites for
IKKphosphorylation, alleviates but does not prevent the NF- activation. This fact suggests

115

that other potential phosphorylation sites co-conduct the signal transduction via NFIdentification and localization of these sites will help in understanding the NF- pathway.
We studied the phosphorylation of IB by GRK5 using HPLC-MS with and without
enrichment (scheme 3.2)

Ib- protein

E.Coli
promoter
gene to be
vector expressed
expression host

Expression

glutathione-S-transferase (GST) tag Lysis and
GST affinity
purification
ESI-MS RP C18 Column Captrap
in solution tryptic digestion
and HPLC-ESI-MS analysis

In vitro phosphorylation
- without enzyme

+ with IKK enzyme
with GRK-5 enzyme

Scheme 3.2. Workflow for the investigation of phosphorylation of IB catalyzed by IKKand
GRK5. The GST-IB protein is expressed in E.Coli and purified with a GST affinity column.
The purified protein undergoes in vitro phosphorylation with no enzyme, with IKKor with
GRK5. Each sample is later digested in solution by trypsin and the tryptic digest is analyzed by
HPLC-ESI-MS. The protein expression and purification was performed by Ms. Wen Qin from
Prof. Narayanan Parameswaran’s group.

116

Maker

-

+

G

Figure 3.12. WB probe of phosphorylation of
104 kD
94 kD

IB using phosphor- IB antibody from
Cell Signaling Technology.

51 kD

-: 0.12 g Ib protein without any enzyme

36 kD
28 kD

+: 0.12 g Ib protein with IKK enzyme

19 kD

G: 0.12 g Ib protein with GRK5 enzyme

After the in vitro phosphorylation, WB analysis confirmed the phosphorylation on IB
by IKKand GRK5 (Figure 3.12). One pmol of digests of GST-IB and phosphorylated GSTIB (by GRK5) were submitted for LC-ESI-MS/MS analysis. Figure 3.13a shows the total ion
current (TIC) chromatogram of the peptides eluted from the HPLC. Each peak may correspond
to one or more peptides with multiple charge states. Figure 3.13b gives the full-MS scan at a
retention time of 18.71 min, and the spectrum contains one dominant peptide signal and tens of
small signals. At a given retention time, the peptides with the top five intensities are
automatically selected for tandem MS spectra. Figure 3.13c reveals the CID-MS/MS spectrum of
the [M+3H]

3+

precursor ion at m/z 768.23, which is a phosphopeptide with two phosphorylation

30

sites ( HDpSGLDpSMKDEEEQMVK). This result confirms that Ser32 and Ser36 are
phosphorylated by GRK5.

117

a

18.71

2

4

6

8

b

10

12

14
16
18
Time (min)

20

22

24

26

28

[M+3H]3+ at m/z 768.23

600

400

800

1000
∆

30H

c

b2

1200
m/z

1400

1600

1800

D pS G L D pS M K D E E Y E Q M V K

y122+

The sequences show cleavage sites for
formation of b and y ions. （∆=-98 Da,

b2 b6∆2+

y4

∆∆
2+
y5 y10

300

500

600

400

700

o=-18 Da and *=-17 Da）
º*
b ∆∆
bº
8

b17
2+ y 2+
y12
15
800

∆∆2+
9
y162+

b17 *2+

900 1000 1100 1200 1300 1400
m/z

Figure 3.13 (a) TIC chromatogram of 1 pmol of a phosphorylated (by GRK5) IB digest (b)
The full-MS scan of peptides eluted at 18.71 min in (a), and (c) CID-MS/MS spectrum of the
[M+3H]

3+

precursor ion at m/z 768.23 in (b).
118

Table 3.2 lists all the peptides identified from 1 pmol of control GST-IB digests, and
no phosphopeptides appear. Table 3.3 lists the peptides identified from the 1 pmol of GST-IB
digests phosphorylated by GRK5. Among those peptides, three phosphopeptides stem from IB
and two phosphopeptides stem from GRK5. Interestingly, the three phosphopeptides from IB
30

have the same sequence ( HDSGLDSMKDEEYEQMVK) but different modifications,
including peptides eluting at 14.77 (pSer 32 and Carbamidomethyl (cam) on Met 37), 18.25
(pSer 32), and 18.71 (pSer 32 and 36) min.

Furthermore signals for their two non-

phosphorylated counterparts also appear at 14.06 and 17.51 min; one has cam on Met 37 and the
other has no modification.
We estimate the phosphorylation efficiency using the ratio of the intensity sum of the
three phosphopeptides to the intensity sum from all five peptides with the sequence
30

HDSGLDSMKDEEYEQMVK. The ratio suggests that 30-min in vitro phosphorylation of

IB by GRK5 (enzyme/substrate mole ratio 1/3) yields ~40 % phosphorylation on
30

HDSGLDSMKDEEYEQMVK. The GRK5 also shows auto-phosphorylation reflected by

DLVDIEQFpSpTVK or DLVDIEQFpSTVK sequences. We found no IB phosphorylation
sites besides Ser 32 and 36, but this could be due to the low stoichiometry of the other
phosphorylations, low sequence coverage from digestion or peptide loss during HPLC.
Phosphopeptide

enrichment

might

enhance

the

phosphopeptide

identification.

Unfortunately, however, using the TiO2-modified membrane enrichment does not reveal any
phosphorylation sites. The presence of SDS or ATP may hinder the enrichment performance, and
methods to remove the 0.4% SDS and 0.25 mM ATP from 0.08 M proteins could lead to loss of
protein. The ATP is essential for phosphorylation, but it may well compete with phosphopeptides
119

for binding sites. Further refinements in methods for ATP and SDS removal may lead to
effective enrichment.
Table 3.2. Peptides identified using HPLC-ESI-MS/MS of 1 pmol of GST-IB digest. The
retention time, peptide mass, sequence and origin protein are listed.
Retention
Time
9.7
9.91
10.55
10.84
11.42
12.18
12.56
12.97
13.05
13.71
13.9
14.08
14.26
14.53
15.52
15.93
16.09
16.76
16.9
17.39
17.6
18.21
18.65
18.84
21.19
21.29
21.72
23.29
23.66
23.92
25.06
25.33

Peptide
Mass
631.62
1661.44
770.86
1089.22
703.72
1012.48
967.84
2868.2
1605.16
666.88
2198.8
752.76
2198.8
2025.92
708.78
1183.02
730.22
1033.06
1027.08
1077.34
2158.06
932.96
1151.86
2754.92
1859.59
2261.29
1574.35
2324.44
2005.76
1442.08
3156.3
2688.55

Peptide sequence
LLDDR
YIADKHNM(cam)LGGC(cam)PK
GLVQPTR
LTQSM(cam)AIIR
GSEPWK
IKGLVQPTR
LEPQEVPR
LLDDRHDSGLDSM(cam)KDEEYEQM(cam)VK
YIADKHNMLGGC(CAR)PK
LVC(cam)FK
HDSGLDSM(cam)KDEEYEQMVK
DEFTLK
HDSGLDSM(cam)KDEEYEQMVK
YEEHLYERDEGDKWR
VDFISK
RIEAIPQIDK
LPEMLK
LTQSMAIIR
IEAIPQIDK
VTYQGYSPY
HDSGLDSM(oxidation)KDEEYEQMVK
ALTMEVIR
M(cam)SPILGYWK
LLDDRHDSGLDSMKDEEYEQMVK
ERAEISM(cam)LEGAVLDIR
VTYQGYSPYQLTWGRPSTR
AEISM(cam)LEGAVLDIR
QQLTEDGDSFLHLAIIHEEK
IAYSKDFETLKVDFLSK
DFETLKVDFLSK
LLLEYLEEKYEEHLYERDEGDKWR
HLAVITNQPEIAEALLGAGC(cam)DPELR
120

Origin
Protein
IB
GST
GST
GST
IB
GST
IB
IB
GST
GST
IB
GST
IB
GST
GST
GST
GST
GST
GST
IB
IB
IB
GST
IB
GST
IB
GST
IB
GST
GST
GST
IB

Table 3.2 (cont’d)
25.69
26.05

1803.13
2270.47

26.27
26.53
26.98
28.16
28.27
29.1
30.03

3865.4
1517.68
2327.77
2358.01
1933.9
2174.38
2230.54

ERAEISMLEGAVLDIR
LLLEYLEEKYEEHLYER
HDSGLDSMC(cam)KDEEYEQM(DE)VK
ELQEIQEIRLEPQEVPR
AEISMLEGAVLDIR
YIAWPLQGWQATFGGGDHPPK
KFELGLEPPNLPYYIDGDVK
GDLAFLNFQNNLQQTPL
TALHLAVDLQNPDLVSLLLK
FELGLEFPNLPYYIDGDVK

GST
GST
IB
GST
GST
GST
IB
IB
GST

Table 3.3. Peptides identified using HPLC-ESI-MS/MS of 1 pmol of GST-IB digest
phosphorylated by GRK5. The retention time, peptide mass, sequence and origin protein are
listed.
Retention
time
10.37
10.63
11.2
12.3
13.82
14.06
14.77
15.27
15.71
15.87
16.31
16.58
16.75
17.21
17.51
18
18.25
18.55
18.71
19.57
21.12

Peptide
mass
771
1090.26
787.56
968.12
753.2
2199.64
2279.6
708.98
1183.28
730.98
1627.24
1033.22
1027.32
916.18
2142.67
933.26
2222.53
1152.42
2302.48
1748.44
2262.01

Peptide sequence
GLVQPTR
LTQSM(cam)AIIR
LPEM(cam)LK
LEPQEVPR
DEFTLK
HDSGLDSM(cam)KDEEYEQMVK
HDpSGLDSM(cam)KDEEYEQMVK
VDFISK
RIEAIPQIDK
LPEMLK
GVNLDHTDDDFYSK
LTQSMAIIR
IEAIPQIDK
ISDLGLAVK
HDSGLDSMKDEEYEQMVK
ALTMEVIR
HDpSGLDSMKDEEYEQMVK
M(cam)SPILGYWK
HDpSGLDpSMKDEEYEQMVK
VGTVGYMAPEVLNNQR
VTYQGYSPYQLTWGRPSTR
121

Origin
protein
GST
GST
GST
IB
GST
IB
IB
GST
GST
GST
GRK5
GST
GST
GRK5
IB
IB
IB
GST
IB
GRK5
IB

Table 3.3 (cont’d)
21.26
22.63
22.71
22.77
22.92
23.06
25.74

3.4

1575.44
1912.54
1554.14
1474.26
1095.22
1150.42
2076.98

AEISM(cam)LEGAVLDIR
DLKPENILLDDYGHIR
DLVDIEQFpSpTVK
DLVDIEQFpSTVK
MSPILGYWK
LLLEYLEEK
SPVFIAQVGQDLVSQTEEK

GST
GRK5
GRK5
GRK5
GST
GST
GRK5

Conclusions
The TiO2-modified membrane effectively facilitated enrichment of the phosphopeptides

from an in vivo phosphorylated fusion protein, Jun-Tev-tau. MS analysis of the enriched eluate
revealed 7 phosphorylation sites on p-tau, three of which seem dominant. Identification of such
sites might aid the design of therapeutic drugs that inhibit phosphorylation.[45-46] In
phosphopeptide enrichment of an IP of p65, we find 8 phosphorylated peptides that correspond
to only 2 unique phosphorylation sites from the p65-associated ribosomal protein. Membranebased enrichment is rapid and facilitates simple identification of phosphopeptides. However,
enrichment of in vitro phosphorylated IB does not reveal any phosphopeptides, perhaps
because ATP or SDS interferes with binding. HPLC-MS, though more time consuming than
membrane-based enrichment and direct MS, is sufficiently sensitive to reveal phosphorylation of
IB at ser32 and ser36. The coupling of on-line membrane enrichment to HPLC-MS is worth
trying in the future, but issues of solvent compatibility, membrane stability (preventing leaching
of TiO2, polyelectrolyte and substrate compounds), and compatibility of enrichment with
compounds in the sample must be addressed. Overall, membrane-based enrichment is attractive
for identifying phosphorylation in modestly complex samples such as fusion proteins.
122

3.5

Acknowledgement
Many thanks to Dr. Dexin Sui and Prof. Min-Hao Kuo for their collaboration in the p-tau

study and Ms. Li Cui for useful discussion, to Dr. Amanda Palumbo, Dr. Teri Lansdell and Prof.
Jetze J. Tepe for their collaboration in the p65 study, to Dr. Xiao Zhou, Ms. Wen Qin, and Prof.
Narayanan Parameswaran for their collaboration in the IBStudy. We are grateful to the U.S.
National Institutes of Health (GM 080511, Bruening and AG039768, Kuo) for funding this work.

123

REFERENCES

124

3.6

References

1.

Aebersold, R.; Mann, M., Mass Spectrometry-Based Proteomics. Nature 2003, 422
(6928), 198-207.

2.

Olsen, J. V.; Blagoev, B.; Gnad, F.; Macek, B.; Kumar, C.; Mortensen, P.; Mann, M.,
Global, in vivo, and Site-Specific Phosphorylation Dynamics in Signaling Networks. Cell
2006, 127 (3), 635-648.

3.

Thingholm, T. E.; Jensen, O. N.; Larsen, M. R., Analytical Strategies for
Phosphoproteomics. Proteomics 2009, 9 (6), 1451-1468.

4.

Witze, E. S.; Old, W. M.; Resing, K. A.; Ahn, N. G., Mapping Protein Post-Translational
Modifications with Mass Spectrometry. Nat. Methods 2007, 4 (10), 798-806.

5.

Han, X.; Aslanian, A.; Yates III, J. R., Mass Spectrometry for Proteomics. Curr. Opin.
Chem. Biol. 2008, 12 (5), 483-490.

6.

Zhou, H.; Low, T. Y.; Hennrich, M. L.; van der Toorn, H.; Schwend, T.; Zou, H.;
Mohammed, S.; Heck, A. J. R., Enhancing the Identification of Phosphopeptides from
Putative Basophilic Kinase Substrates Using Ti (IV) Based IMAC Enrichment. Mol. Cell.
Proteomics 2011, 10 (10).

7.

Leitner, A., Phosphopeptide Enrichment Using Metal Oxide Affinity Chromatography.
Trac-Trend Anal. Chem. 2010, 29 (2), 177-185.

8.

Sergeant, N.; Delacourte, A.; Buee, L., Tau Protein as a Differential Biomarker of
Tauopathies. Biochim. Biophys. Acta, Mol. Basis Dis. 2005, 1739 (2-3), 179-197.

9.

Wang, J. Z.; Wu, Q. L.; Smith, A.; Grundke-Iqbal, I.; Iqbal, K., Tau Is Phosphorylated by
GSK-3 at Several Sites Found in Alzheimer Disease and Its Biological Activity Markedly
Inhibited Only after It Is Prephosphorylated by a-Kinase. FEBS Lett. 1998, 436 (1), 28-34.

10.

Gilmore, T. D., Introduction to NF-[Kappa]B: Players, Pathways, Perspectives.
Oncogene 2006, 25 (51), 6680-6684.

125

11.

Brasier, A., The NF-Κb Regulatory Network. Cardiovasc. Toxicol. 2006, 6 (2), 111-130.

12.

Perkins, N. D., Integrating Cell-Signalling Pathways with NF-[Kappa]B and IKK
Function. Nat Rev Mol Cell Biol 2007, 8 (1), 49-62.

13.

Meffert, M. K.; Chang, J. M.; Wiltgen, B. J.; Fanselow, M. S.; Baltimore, D., NF[Kappa]B Functions in Synaptic Signaling and Behavior. Nat. Neurosci. 2003, 6 (10),
1072-1078.

14.

Tian, B.; Brasier, A. R., Identification of a Nuclear Factor Kappa B-Dependent Gene
Network. Recent Prog. Horm. Res. 2003, 58 (1), 95-130.

15.

Albensi, B. C.; Mattson, M. P., Evidence for the Involvement of TNF and NF-Κb in
Hippocampal Synaptic Plasticity. Synapse 2000, 35 (2), 151-159.

16.

Vallabhapurapu, S.; Karin, M., Regulation and Function of NF-Κb Transcription Factors
in the Immune System. Annu. Rev. Immunol. 2009, 27 (1), 693-733.

17.

Patial, S.; Luo, J.; Porter, K. J.; Benovic, J. L.; Parameswaran, N., G-Protein-CoupledReceptor Kinases Mediate TNFα-Induced NF-Κb Signalling Via Direct Interaction with
and Phosphorylation of IκBα. Biochem. J 2009, 425 (1), 169-178.

18.

Patial, S.; Shahi, S.; Saini, Y.; Lee, T.; Packiriswamy, N.; Appledorn, D. M.; LaPres, J. J.;
Amalfitano, A.; Parameswaran, N., G-Protein Coupled Receptor Kinase 5 Mediates
Lipopolysaccharide-Induced NFκB Activation in Primary Macrophages and Modulates
Inflammation in vivo in Mice. J. Cell. Physiol. 2011, 226 (5), 1323-1333.

19.

Wang, W.-H.; Palumbo, A. M.; Tan, Y.-J.; Reid, G. E.; Tepe, J. J.; Bruening, M. L.,
Identification of P65-Associated Phosphoproteins by Mass Spectrometry after on-Plate
Phosphopeptide Enrichment Using Polymer-Oxotitanium Films. J. Proteome Res. 2010,
9 (6), 3005-3015.

20.

Wagner, U.; Utton, M.; Gallo, J. M.; Miller, C. C. J., Cellular Phosphorylation of Tau by
GSK-3 Beta Influences Tau Binding to Microtubules and Microtubule Organisation. J.
Cell Sci. 1996, 109, 1537-1543.

21.

Hanger, D. P.; Noble, W., Functional Implications of Glycogen Synthase Kinase-3Mediated Tau Phosphorylation. Int. J. Alzheimers Dis. 2011, 2011, 352805.

126

22.

Rankin, C.; Sun, Q.; Gamblin, T., Tau Phosphorylation by GSK-3&#946; Promotes
Tangle-Like Filament Morphology. BioMed Central: 2007.

23.

Smet-Nocca, C.; Broncel, M.; Wieruszeski, J.-M.; Tokarski, C.; Hanoulle, X.; Leroy, A.;
Landrieu, I.; Rolando, C.; Lippens, G.; Hackenberger, C. P. R., Identification of OGlcnac Sites within Peptides of the Tau Protein and Their Impact on Phosphorylation.
Mol. Biosyst. 2011, 7 (5), 1420-1429.

24.

Palumbo, A. M.; Reid, G. E., Evaluation of Gas-Phase Rearrangement and Competing
Fragmentation Reactions on Protein Phosphorylation Site Assignment Using Collision
Induced Dissociation-MS/MS and MS(3). Anal. Chem. 2008, 80 (24), 9735-9747.

25.

Palumbo, A. M.; Smith, S. A.; Kalcic, C. L.; Dantus, M.; Stemmer, P. M.; Reid, G. E.,
Tandem Mass Spectrometry Strategies for Phosphoproteome Analysis. Mass Spectrom.
Rev. 2011, 30 (4), 600-625.

26.

Kim, M.-S.; Zhong, J.; Kandasamy, K.; Delanghe, B.; Pandey, A., Systematic Evaluation
of Alternating Cid and Etd Fragmentation for Phosphorylated Peptides. Proteomics 2011,
11 (12), 2568-2572.

27.

Hanger, D. P.; Anderton, B. H.; Noble, W., Tau Phosphorylation: The Therapeutic
Challenge for Neurodegenerative Disease. Trends Mol. Med. 2009, 15 (3), 112-119.

28.

Thingholm, T. E.; Jensen, O. N.; Robinson, P. J.; Larsen, M. R., Simac (Sequential
Elution from IMAC), a Phosphoproteomics Strategy for the Rapid Separation of
Monophosphorylated from Multiply Phosphorylated Peptides. Mol. Cell. Proteomics
2008, 7 (4), 661-671.

29.

Reynolds, C. H.; Betts, J. C.; Blackstock, W. P.; Nebreda, A. R.; Anderton, B. H.,
Phosphorylation Sites on Tau Identified by Nanoelectrospray Mass Spectrometry. J.
Neurochem. 2000, 74 (4), 1587-1595.

30.

Cohen, P.; Frame, S., The Renaissance of GSK3. Nat Rev Mol Cell Biol 2001, 2 (10),
769-776.

31.

Hoffmann, A.; Baltimore, D., Circuitry of Nuclear Factor Kappa B Signaling. Immunol.
Rev. 2006, 210, 171-186.

127

32.

Karin, M.; Lin, A., NF-Kappa B at the Crossroads of Life and Death. Nat. Immunol. 2002,
3 (3), 221-227.

33.

Sen, R.; Baltimore, D., Multiple Nuclear Factors Interact with the Immunoglobulin
Enhancer Sequences. Cell 1986, 46 (5), 705-716.

34.

Vallabhapurapu, S.; Karin, M., Regulation and Function of NF-Kappa B Transcription
Factors in the Immune System. Annu. Rev. Immunol. 2009, 27, 693-733.

35.

Bonizzi, G.; Karin, M., The Two NF-Kappa B Activation Pathways and Their Role in
Innate and Adaptive Immunity. Trends Immunol. 2004, 25 (6), 280-288.

36.

O'Shea, J. M.; Perkins, N. D., Regulation of the Reia (P65) Transactivation Domain.
Biochem. Soc. Trans. 2008, 36, 603-608.

37.

Perkins, N. D., Post-Translational Modifications Regulating the Activity and Function of
the Nuclear Factor Kappa B Pathway. Oncogene 2006, 25 (51), 6717-6730.

38.

Mann, M.; Ong, S. E.; Gronborg, M.; Steen, H.; Jensen, O. N.; Pandey, A., Analysis of
Protein Phosphorylation Using Mass Spectrometry: Deciphering the Phosphoproteome.
Trends Biotechnol. 2002, 20 (6), 261-268.

39.

Ryo, A.; Suizu, F.; Yoshida, Y.; Perrem, K.; Liou, Y. C.; Wulf, G.; Rottapel, R.;
Yamaoka, S.; Lu, K. P., Regulation of NF-Kappa B Signaling by Pin1-Dependent Prolyl
Isomerization and Ubiquitin-Mediated Proteolysis of P65/Rela. Mol. Cell 2003, 12 (6),
1413-1426.

40.

Lapko, V. N.; Smith, D. L.; Smith, J. B., Identification of an Artifact in the Mass
Spectrometry of Proteins Derivatized with Iodoacetamide. J. Mass Spectrom. 2000, 35
(4), 572-575.

41.

Kruger, R.; Hung, C. W.; Edelson-Averbukh, M.; Lehmann, W. D., IodoacetamideAlkylated Methionine Can Mimic Neutral Loss of Phosphoric Acid from
Phosphopeptides as Exemplified by Nano-Electrospray Ionization Quadrupole Time-ofFlight Parent Ion Scanning. Rapid Commun. Mass Spectrom. 2005, 19 (12), 1709-1716.

42.

Reid, G. E.; Roberts, K. D.; Simpson, R. J.; O'Hair, R. A. J., Selective Identification and
Quantitative Analysis of Methionine Containing Peptides by Charge Derivatization and
Tandem Mass Spectrometry. J. Am. Soc. Mass Spectrom. 2005, 16 (7), 1131-1150.
128

43.

Wool, I. G., Extraribosomal Functions of Ribosomal Proteins. Trends Biochem. Sci 1996,
21 (5), 164-165.

44.

Lindstrom, M. S., Emerging Functions of Ribosomal Proteins in Gene-Specific
Transcription and Translation. Biochem. Biophys. Res. Commun. 2009, 379 (2), 167-170.

45.

Hong, M.; Chen, D. C. R.; Klein, P. S.; Lee, V. M. Y., Lithium Reduces Tau
Phosphorylation by Inhibition of Glycogen Synthase Kinase-3. J. Biol. Chem. 1997, 272
(40), 25326-25332.

46.

Mazanetz, M. P.; Fischer, P. M., Untangling Tau Hyperphosphorylation in Drug Design
for Neurodegenerative Diseases. Nat. Rev. Drug Discovery 2007, 6 (6), 464-479.

129

Chapter 4. Limited Proteolysis Via msec-Digestions in
Protease-Modified Membranes
(This chapter was submitted for publication as part of an article in Analytical Chemistry)
Sequential adsorption of poly(styrene sulfonate) (PSS) and proteases in porous nylon
yields

enzymatic

membrane

reactors

for

limited

protein

digestion.

3

Although a high local enzyme density (~30 mg/cm ) and small pore diameters in the membrane
lead to digestion in < 1 sec, the low membrane thickness (170 m) affords control over residence
times at the ms level to limit digestion. Apomyoglobin digestion demonstrates that peptide
lengths increase as the residence time in the membrane decreases. Moreover, electron transfer
dissociation (ETD) tandem mass spectrometry (MS/MS) on a large myoglobin proteolytic
peptide (8 kD) provides a resolution of 1-2 amino acids. Under denaturing conditions, limited
membrane digestion of bovine serum albumin (BSA) and subsequent ESI-Orbitrap MS analysis
reveal large peptides (3-10 kD) that increase the sequence coverage from 53 % (2-s digestion) to
82 % (0.05-s digestion).

4.1

Introduction
Proteolysis is a vital initial step in most analyses of proteins by MS. Traditional in-

solution digestion employs a dilute protease solution to avoid sample contamination and selfdigestion of the enzyme, but the low protease concentration leads to extended digestion times.
To increase throughput and facilitate online MS analysis, a number of research groups now
employ immobilized enzymes for proteolysis.[1-3] Immobilization greatly reduces self-digestion
3

even at high local enzyme density (e.g. 30 mg/cm ),[4-6] and such high densities allow protein
130

digestion in as little as a few seconds. In addition, adsorption stabilizes the enzyme in many
cases.[1] Protease immobilization can occur via covalent,[7] hydrophobic[4, 6] or electrostatic
interactions,[8] and supports for these enzymes include resins,[9] monoliths,[5] polymeric
microfluidic channels[4, 10] and membranes.[6] All of these substrates provide relatively high
surface areas for protease binding. However, porous membranes are unique in that their minimal
thickness (typically 10-170 m) should afford fine control over proteolysis time in flow-through
digestion. When coupled with simple syringe pumps, membranes provide digestion times
ranging from a few milliseconds to several seconds.
This chapter exploits short residence times in enzyme-modified membranes to perform
limited proteolysis of BSA and apomyoglobin. We hypothesize that decreasing membrane
residence times will lead to longer peptides with more charge (Figure 4.1), and this is
particularly the case for pepsin-modified membranes. Unlike solution digestion, where partially
digested peptides continuously compete with intact proteins for enzyme sites that catalyze
additional proteolysis, convective flow in membranes rapidly separates digested peptides from
immobilized enzymes. Large proteolytic peptides often contain more basic residues than small
tryptic peptides and are particularly amenable to ETD MS/MS analyses with high amino acid
resolution, which is important for studies of hydrogen/deuterium exchange (HDX) to determine
amino acid accessibility.[11-12] Current methods for creating large proteolytic peptides employ
expensive enzymes such as Asp-N and Glu-C that cut proteins only before or after one type of
amino acid.[13] However, these methods work only for a few proteins that contain a limited
number of cleavable sites. Limited digestion in membranes should be applicable to a wide range
of proteins because generation of long peptides relies on rapid convection of partially digested
peptides away from the protease.

131

Figure 4.1. Cartoon of controlled digestion using a membrane containing immobilized pepsin
and a miniaturized holder with a syringe pump. (The apomyoglobin structure was generated
from PDB 1MBN.）

132

4.2

Experimental

4.2.1 Materials
Pepsin from porcine gastric mucosa (lyophilized powder, 3200-4500 units/mg protein),
trypsin from bovine pancreas (type I, 12200 units/mg solid), -chymotrypsin from bovine
pancreas (L-1-tosylamide-2-phenylethyl chloromethyl ketone treated to inactivate residual
trypsin activity, type VII, salt-free lyophilized power), apomyoglobin (protein sequencing grade
from horse skeletal muscle, salt-free lyophilized powder), PSS (average molecular weight
~70,000), and bovine serum albumin (BSA) were purchased from Sigma-Aldrich. Important
chemicals for digestion and analysis included Tris-HCl (Invitrogen), urea (J.T.Baker), 1,4-dithioDL-threitol (DTT) (BioChemika), iodoacetamide (Sigma), ammonium bicarbonate (Columbus
Chemical),

2,5-dihydroxybenzoic

acid

(DHB,

Sigma),

tris(2-carboxyethyl)

phosphine

hydrochloride (TCEP, >98%, Fluka), ProgentaTM non-ionic acid labile surfactant (NALS,
Protabio), and formic acid (FA, > 96%, Spectrum) .

4.2.2 Membrane and holder
The unmodified nylon membranes (25 mm-diameter, 0.45 m nominal pore size) were
obtained from Millipore (HNWP02500). Membrane modification employed an Amicon cell
2

(model 8010, Millipore) that exposed a membrane area of 3.8 cm . A peristaltic pump pulled
solutions through the membrane. For experiments with small sample volumes (<1 mL), a 6 mmdiameter piece of membrane was placed in a flangeless fitting system (A-424, Upchurch
Scientific) connected to 1/16 inch OD tubing via ferrules. The membrane was supported by a frit
2

in the holder. The membrane area exposed to sample was 0.02 cm , and sample injection
133

occurred via a syringe pump (Cole-Parmer 74900 series) using a 1-mL or 5-mL disposable
syringe.

4.2.3 Modification of membranes with proteases
Modification of UV/ozone-cleaned (10 min) nylon membranes with trypsin was reported
previously, but I increased the trypsin concentration in the adsorption solution from 0.6 to 1.0
mg/mL.[8] For modification with pepsin, the procedure was similar and also occurred through
consecutive adsorption of PSS and proteases. Ten mL of a 0.02 M PSS, 0.5 M NaCl solution
(pH=2.3) was first passed through the membrane at 2 mL/min, followed by 30 mL of deionized
water at the same flow rate. Subsequently, 4 mL of 2 mg/mL pepsin in 5% v/v FA was circulated
through the membrane at 1 mL/min for one hour. After protease deposition, the membrane was
rinsed with 30 mL of 5% v/v FA. Modified membranes were dried and stored in a desiccator.
Using a Bradford assay with calibration curves, the amount of pepsin in the membrane was
determined from the decrease in pepsin concentration in the loading solution after circulation
through the membrane. The amounts of trypsin immobilized in membranes were similarly
determined from direct UV absorbance at 280 nm using a Nanodrop

TM

UV/Vis Spectrometer

(Thermo).

4.2.4 Controlled apomyoglobin digestion with a pepsin-modified membrane
Without urea denaturation or reduction, 100 g of apomyoglobin was dissolved in 1 mL
of 5% v/v FA, and this solution was passed through a small membrane (Figure 4.1) at flow rates
from 0.1 to 30 mL/h. We calculated the residence time by dividing the membrane pore volume
2

(estimated assuming 50% porosity, a 170-m thickness, and for the small holder 0.02 cm of
134

external area) by the flow rate. Consequently, the residence times for 0.1 and 30 mL/h flow rates
were 6 s and 0.02 s, respectively. FiftyL of each digest was collected, dried with a Speed-Vac
and reconstituted in 50 L of 1% acetic acid/49% H2O/50% methanol. The same experiments
were also carried out with a trypsin-modified membrane, except that the apomyoglobin buffer
was 10 mM NH4HCO3. For in-solution peptic digestion, 100 g of apomyoglobin in 200 L 5%
v/v FA was mixed with 5 g of pepsin and incubated at 37 º for 5 min or 20 s. Two hundred L
C
of acetonitrile was added to stop the reaction, and the solution was immediately frozen with
liquid nitrogen and lyophilized with a Speed-Vac.

4.2.5 Controlled BSA digestion with a pepsin-modified membrane
BSA was digested with urea denaturation and TCEP reduction. Thus, 100 g of BSA was
dissolved in 10 L of 6 M urea and mixed with 1 L of 0.5 M TCEP for 30 min at 56 º To
C.
make a 0.1 mg/mL BSA solution, 989 L of 5% v/v FA was added to the above 11 L samples.
This solution was passed through a small membrane at 0.3, 1.5, 3, 6 and 12 mL/h (the 5-mL
syringe was used for the last two flow rates). Two hundred L of each digest was collected,
dried with a Speed-Vac and reconstituted in 200 L of 1% acetic acid/49% H2O/50% methanol.
A similar experiment occurred with a trypsin-modified membrane, except that we applied a
NALS as the denaturation agent following the manufacture’s protocol with slight modification:
100 g of BSA was dissolved in 20 L of 0.1 % NALS at pH 7.8 (50 mM NH4HCO3), reduced
with 5 L of 10 mM DTT for 30 min at 56 º and allowed to react with 10 L of 100 mM
C
iodoacetamide in the dark for 30 min.

135

4.2.6 Mass spectrometry and data analysis
An LTQ XL ion trap mass spectrometer equipped with a vMALDI source (Thermo
Fisher) was used to perform MALDI-MS and CID-MS/MS analyses. Typically, 1 L of sample
solution was spotted onto the stainless steel MALDI plate, followed by addition of 1 L of
matrix solution (40 mg/mL of 2,5 DHB in 1% TFA, 49% H2O, 50% ACN) and drying. MALDIMS spectra were acquired using a laser power of 10 µ across the m/z range 300-2000 (normal
J
mass range) or 800-4000 (high mass range). Spectra were the average of 30-100 scans.
For ESI-MS analyses, 40 µL of samples in 1% acetic acid, 49% H2O and 50% methanol
were loaded into a Whatman multichem 96-well plate (Sigma Aldrich) and sealed with Teflon
Ultra Thin Sealing Tape (Analytical Sales and Services, Prompton Plains, NJ). Then, using an
Advion Triversa Nanomate nano-electrospray ionization (nESI) source (Advion, Ithaca, NY)
with a spray voltage of 1.4 kV and a gas pressure of 1.0 psi, samples were introduced into a high
resolution accurate mass LTQ Orbitrap Velos mass spectrometer (Thermo Fisher) equipped with
a dual pressure ion trap, Higher-Energy Collisional Dissociation (HCD) cell and ETD. The ion
source interface settings (inlet temperature of 200 oC and S-Lens value of 60 %) were optimized
to maximize the sensitivity of the precursor ions while minimizing ‘in-source’ fragmentation.
High resolution mass spectra, as well as all ETD-MS/MS spectra, were acquired in positive
ionization mode using the FT analyzer operating at 100,000 resolving power. Spectra were
typically signal averaged for 30-300 scans, and peptide precursor ions were mass-selected using
an isolation width of 5 Da. In ETD-MS/MS, fragmentation of isolated ions was performed when
4

an automatic gain control (AGC) target value of 5.0 × 10 was reached within a 100 ms
maximum injection time in the ion trap. The AGC value and maximum injection time of the

136

5

reagent ion (fluoranthene anion) were set as 1.0 × 10 and 100 ms, respectively, and the reaction
time was optimized as 15 ms for the +10 charge state.
Peptides in Tables 4.1 and 4.2 were identified using peptide mass fingerprinting. If the
monoisotopic peak (m/z=a) could be found, then the monoisotopic peptide mass was calculated
by using the equation, M+H= (a*z-(z-1)*1.0078). The calculated mass was matched with a mass
from in silico digestion of the target protein with ProteinProspector (http://prospector.ucsf.edu).
For ESI-Orbitrap data, the mass tolerance was 5 ppm. If the monoisotopic peak could not be
located, the mass of the highest peak served as an estimate of the average mass of the target
peptide. The simulated spectrum of the potential peptide match was then compared with the
experimental spectrum, and the peptide was assigned if specific isotopic signals in the two
spectra matched within 5 ppm. For MALDI-LTQ data, the mass tolerance was 100 ppm.

4.3

Results and discussion

4.3.1 Membrane modification with protease
We previously showed that alternating adsorption of PSS and trypsin effectively
immobilizes trypsin in nylon membranes.[8] In 2.7 mM HCl, the protonated trypsin (isoelectric
point, pI, of 10.2) adsorbs to negatively charged PSS.[8, 14-15] Pepsin has a much lower pI (2.8),
so we both adsorb this protease and perform peptic digestion using solutions containing 5% v/v
FA (pH=2).

Based on analyses of protease adsorption solutions, the protease/PSS-modified
3

3

nylon membranes contain ~30 mg of trypsin/cm of pores, and ~60 mg pepsin/cm (Figure 4.2).
These values assume a membrane porosity of 50%. Control experiments show that pepsin and
trypsin do not adsorb to the bare membrane without PSS. The high local protease density in

137

3

modified membranes, compared to 0.025 mg/cm for in-solution digestion, leads to membranebased protein digestion in times ranging from milliseconds to seconds, depending on the protein.

Bradford dye absorbance

0.3
0.25

R2=0.999

0.2
0.15

Original pepsin solution

0.1

Pepsin solution after modification

0.05
0

0

0.2

0.4

0.6

0.8

1

1.2

Concentration of pepsin in solution/ (mg/mL)
Figure 4.2. Bradford assay calibration curve for pepsin, and the absorbance of a pepsin solution
(0.4 mg/mL after a 5-fold dilution) before and after circulating through the membrane. By
subtraction, we estimate that 1.8 mg of pepsin adsorbed to the PSS-modified nylon membrane.
3

This corresponds to a local density of 60 mg pepsin per cm of pore.

4.3.2 Apomyoglobin digestion with different membrane residence times
Due to its simple secondary structure and high purity, apomyoglobin served as the first
model protein to test our hypothesis that decreasing the membrane residence time will give
longer proteolytic peptides. Figure 4.3 shows manually deconvoluted ESI-Orbitrap mass spectra

138

of myoglobin digested by passing solutions through pepsin-modified membranes at flow rates of
0.1, 1, 10 and 30 mL/h, which correspond to residence times of 6, 0.6, 0.06 and 0.02 s,
respectively. As the residence time decreases, large peptides emerge and the strongest signals
shift to a higher m/z region. (Figure 4.4 shows the original spectra, and Table 4.1 lists peptides
identified from the spectra along with their monoisotopic masses, amino acid positions and
cleavage sites). With the 6-s residence time, all detected peptides have masses less than 5 kD,
suggesting a relatively complete digestion. In-solution digestion for 5 min or 20 s and membrane
digestion for 6 s give similar mass spectra that yield 100 % sequence coverage with a minimum
of 5 peptides (2, 7, 8, 11 and 13) (Figure 4.5). For the 0.6 and 0.06-s membrane-based digestions,
covering the entire 153 amino acid sequence requires 4 peptides (2, 7, 8, and 16) and only 3
peptides (7, 15 and 16), respectively. As the residence time decreases, the signal for peptide 16
replaces the signals for peptides 11 and 13 due to missed cleavage. Similarly, signals for
peptides 2 and 8 decrease in absolute intensity and the signal for peptide 15 (the combination of
peptides 2 and 8) grows. For the 0.02-s digestion, signals for the intact protein (20) and peptide
19 (amino acids 30-153) dominate the spectrum, indicating incomplete digestion.

139

Figure

4.3.

Manually

deconvoluted ESI-Orbitrap mass
spectra of 0.1 mg/mL (6 M)
apomyoglobin

digested

with

residence times of (a) 6, (b) 0.6,
(c) 0.06 and (d) 0.02 s in a pepsinmodified membrane. Signals with
intensities

above 1% of the

highest signal are assigned to the
specific peptides if possible and
converted to the m/z value for the
+1 charge state. The normalized
intensities in the spectra are the
sum of signal intensities for all
detected charge states of a given
peptide. The inset table lists all
peptides identified in the four
spectra (AA represents the amino
acid sequences).

140

11

a
11

2

13
13
6
12
9 5

1 10

b

811
4 12

13 3

7

16 11 12 16 13 3

7

11

13

11

13

2

8
12

56

c

15

7

2

16

16
16
13
15

15
11

16
13

8

16
14 19
15
18

11

11

d

19
19

15

20
20

19 16
13

2

500 600

700

800

20 19

20

19
19

17
16
19 18
18

16

19

20
19

20
19

7

20

15

20
18

19
20

900 1000 1100 1200 1300 1400 1500 1600
m/z

Figure 4.4. ESI-Orbitrap mass spectra of 0.1 mg/mL (6 M) apomyoglobin digested by a pepsinmodified membrane in (a) 6, (b) 0.6, (c) 0.06 and (d) 0.02 s. Labels refer to the peptide list in
Table 4.1

141

11

a
2

11

13

13
10

6

1

2

8

11

9 5 12

4

13

12

3

7

11

b

13

11

2
6

10

11
2

12

13
12

1

8

3

13

11

c

13

4

8

2

11

13

13

7

11
3

2

500

600

12

6

700

800
m/z

3

900

1000

1100

Figure 4.5. ESI-Orbitrap mass spectra of (a) 0.1 mg/mL (6 M) apomyoglobin digested by a
pepsin-modified membrane for 6 s and (b, c) 100 g apomyoglobin digested by 5 g pepsin in
0.2 mL of 5% v/v FA. Digestion times are 5 min and 20 s for (b) and (c), respectively. Both
solution digests are diluted to 0.1 mg/mL (6 M) before MS analysis. The labels refer to the
peptide list in Table 4.1.
142

Table 4.1. Apomyoglobin peptides identified from ESI-Orbitrap mass spectra of protein digested
in a pepsin-modified membrane. Peak number, monoisotopic mass, amino acids, cleavage sites
and missed cleavage sites are listed.
Peak
Number

Peptide
Monoisotopic Mass

Amino
Acids

Cleavage
Sites

Missed Cleavage Sites

1
2

1802.03
1856.97

70-86
138-153

L, L
L, End

2A, 2L, I
2A, I, L,Y, 2F

3
4

1921.98
2501.27

12-29
111-134

L, L
A, A

2A, 2L
4A, 2I, L

5
6
7
8

2856.48
2887.45
3134.55
3242.67

111-137
107-134
1-29
107-137

A, L
F, A
L
F, L

4A, 3L, 2I
5A, 3I, L
2A, 2L, I, 2W
5A, 3I, 3L

9

3506.97

77-106

L, F

3A, 3L, 2I, Y

10
11

3743.16
4132.35

70-103
70-106

L, Y
L, F

4A, 4L, 3I
5L, 4A, 3I, Y

12
13
14
15

4269.24
4651.51
4694.43
5080.61

33-69
30-69
111-153
107-153

L, L
L, L
A, End
F, End

3F, 3L, 2A
4L, 3F, 2A, I
7A, 3I, 3L, 3F, Y
6A, 4L, 3I, 3F, Y

16
17

8764.84
8863.86

30-106
62-142

L, F
L, I

10L, 6A, 4I, 3F, Y
9L, 9A, 6I, 3F, Y

18

11988.49

30-137

L, L

12L, 11A,7I, 5F, Y

19
20

13826.44
16941.97

30-153
1-153

L, End
End

14L, 13A, 8I, 7F, 2Y
17L, 15A, 9I, 7F, 2Y, 2W

Apomyoglobin Sequence:
1

GLSDGEWQQVLNVWGKVEADIAGHGQEVLIRLFTGHPETLEKFDKFKHLKTEAE
MKASEDLKKHGTVVLTALGGILKKKGHHEAELKPLAQSHATKHKIPIKYLEFISDA
IIHVLHSKHPGDFGADAQGAMTKALELFRNDIAAKYKELGFQG

143

We also explored residence time-dependent digestion of apomyoglobin using trypsin/PSS
-modified membranes and observed large tryptic peptides at faster flow rates. However, a high
level of intact apomyoglobin co-exists with these peptides (data not shown). The presence of
intact protein confirms that convection rapidly removes protein from the membrane, in contrast
to a previous study that employed trapping in nanopores to enhance digestion.[16]

In

conventional in-solution digestion with several proteases, apomyoglobin undergoes slow
proteolysis at pH 7-8.[17] Apomyoglobin denaturation at pH 2 might explain why at low
residence times peptic digestion is more efficient than tryptic digestion for this protein. Thus,
pepsin is superior to trypsin for creating large proteolytic peptides from apomyoglobin.

4.3.3 ESI-Orbitrap ETD-MS/MS analysis of a large apomyoglobin peptide
Figure 4.6 summarizes the results of apomyoglobin digestion in pepsin-modified
membranes in terms of number of peptides, average peptide length and average charge state.
Overall, shorter residence times generate larger peptides with more charge. ETD analysis of
peptide sequences requires multiple charges to generate abundant fragments,[18] and Coon and
coworkers suggest that a low residues/charge ratio and three or more charges are the ideal
conditions for effective ETD MS/MS.[19] Short membrane digestion (0.06 s) gives a myoglobin
peptide (16, amino acids 30-106) with a mass of 8764.93 Da and a +10 charge state, and Figure
4.7 shows the ETD MS/MS spectrum for this peptide. We identified 46 c ions and 52 z ions,
which represent cleavage of 61 out of 76 total amino acid junctions. The average spatial
resolution is less than two amino acid residues, equivalent to the results carried out by more
complex instrumentation in Top-Down FT-ICR ECD analysis.[20]

144

Number of peptides
Average length of peptides
Average charge of peptides at most abundant peak

66

58

34

28
13
4.3
61

10

11

5

2
3
0.6
0.06
Residence time / Second

7.8

8.6

8
4
0.02

Figure 4.6. Number (blue), average length (red), and average charge of peptides at the most
abundant peak (green) in ESI-Orbitrap mass spectra of 0.1 mg/mL apomyoglobin digested in a
membrane for 6 s, 0.6 s, 0.06 s and 0.02 s.

145

c3

30I

RLFTGHPETLEKFDKFKHLKTEAEM
K A S E D L K K H G T V V L T A LaG G I L K K K G H
HEAELKPLAQSHATKHKIPIKYLEF
70

c6

z132+
c

c5

14

z102+
c92+ c112+z
5
2+ c4
c3 c8

c183+

z5

z142+
2+

z294+

[M+8H]8+

[M+9H]9+

z475+ 4+
c25 c546+ c35 3+ c364+
c26
3+
4+
2+
c34
z182+ c z28
c17
z172+
9
z374+
4+
3+

c182+

c33

450 550 650 750 850 970 990 1010 1030 1050 1070 1090

400

600

800

1000 1200 1400 1600 1800
m/z

Figure 4.7. ESI-Orbitrap ETD-MS/MS spectrum of the [M+10H]

10+

precursor ion at 878.00.

The sequence shows cleavages site for formation of major c and z ions.

This high spatial resolution coupled with rapid, acidic peptic digestion may be
particularly attractive for HDX studies that provide structural information.[11-12] Short, acidic
digestion should reduce either back-exchange or over-exchange in HDX, and high spatial
resolution will increase the confidence in exchange site localization. Additionally, acidic
digestion might facilitate characterization of membrane proteins that are insoluble at neutral pH.
146

4.3.4 In-membrane BSA digestion under denaturing conditions
Sequence coverage is a critical parameter for MS-based protein characterization,
particularly for comprehensive examination of posttranslational modifications. With a molecular
mass of 66 kD (583 amino acids) and 17 disulfide bonds, BSA is a reasonable model for
examining how membrane-based digestion time affects sequence coverage for a medium-sized,
denatured protein. The 17 disulfide bonds make denaturation and reduction prerequisites for
membrane digestion.
Figure 4.8 shows the ESI-Orbitrap MS spectra (m/z 540-1440) of BSA digested in
membranes using flow rates of 12 and 0.3 mL/h, or residence times of 0.05 and 2 s, respectively.
The 0.05-s residence time represents limited digestion, and the 2-s residence time usually
provides nearly complete digestion. Peak labels indicate identified BSA peptides in order of
+

increasing mass of (M+H) from 628.414 for peptide 1 to 9204.38 for peptide 52. Comparing
the 2-s (Figure 4.8a) and 0.05-s (Figure 4.8b) digestions, the shorter time reveals more peptides
at a high m/z range.

Figure 4.9 presents the peptide size distributions in the two digestions.

Remarkably, short digestion yields 10 peptides with masses >5 kD, whereas the 2-s digestion
shows only 1 peptide with a mass greater than 3 kD. With 0.05-s digestion, 12 peptides (17, 21,
33, 35, 39, 44, 45, 46, 48, 50, 51 and 52) cover 82 % of the 583 amino acids, while the 2-s
membrane digestion needs 19 peptides (1, 2, 8, 9, 10, 11, 17, 18, 20, 22, 26, 27, 28, 29, 30, 31,
32, 33 and 36) to cover just 53 % of the sequence. (The inclusion of other identified peptides
does not increase sequence coverage, and we included only peptides with signals >1.5% of the
highest signal in the spectrum.) The low ionization efficiency of some small peptides may
contribute to the low sequence coverage with the 2-s digestion time, whereas longer peptides

147

with more basic residues (Lys and Arg) provide higher ionization efficiency in positive-ion
mode.[21]
22

a
7

2
1

30

22

17

8
10 33
9
11

5 6
20
36
31
32
18 29

33
27
28

26
33

b
22

33

22

17

51 44

38
1

21

35

50

45 47

48

600

700

800

900

1000

m/z

1100

38

1200

44
52

1300

33

1400

Figure 4.8. ESI-Orbitrap mass spectra of 0.1 mg/mL BSA (1.5 M in 0.06 M urea) digested in a
pepsin-modified membrane using (a) 2-s and (b) 0.05-s residence times. The numbers refer to the
identified BSA peptides where m/z values increase with the peptide number. (Table 4.2 lists all
peptides along with their monoisotopic masses and sequences.)

148

2-s digestion

0.05-s digestion

25
16
8

7

6
2

2

1

Mass of (M+H)+
Figure 4.9. Peptides size distributions in ESI-Orbitrap mass spectra of 0.1 mg/mL BSA digested
in a pepsin-modified membrane for 2-s (blue, 32 peptides) and 0.05-s (red, 35 peptides).
Table 4.2. BSA peptides identified from ESI-Orbitrap mass spectra of protein digested in a
pepsin-modified membrane. Peak number, monoisotopic mass, amino acids, cleavage sites and
missed cleavage sites are listed.
Peak
Number

Peptide
Monoisotopic Mass

Amino
Acids

Cleavage
Site

Missed Cleavage
Sites

1

628.414

455-459

L, L

I, L

2

688.424

228-233

F, L

NONE

3

713.419

568-574

F, L

A

4

731.42

213-218

A, L

W, A

5

868.423

544-550

L, F

NONE

6

889.499

573-583

L, END

A, L

7

1081.57

219-227

L, F

A, F

8

1306.74

346-356

L, L

2A, 2L, Y

9

1351.67

154-164

L, F

3Y, L, A

10

1368.68

395-406

F, L

L, Y, F, A

11

1414.81

179-190

L, A

I, L

12

1421.81

532-543

L, L

L, A

13

1499.84

516-528

L, L

I, A

14

1517.6

554-567

F, F

3A

15

1538.77

488-501

F, F

2A, L, Y

16

1548.84

374-386

F, L

2L

149

Table 4.2 (cont’d)
17

1583.9

568-583

F, END

2A, 2L

18

1632.86

142-154

I, L

2A, 2Y, L, F

19

1763.01

529-543

L, L

2L, A

20

1793.97

213-227

A, F

2A, L, F, W

21

1834.78

551-567

F, F

4A, F

22

1858.09

407-422

L, L

I, Y

23

1868.93

331-345

L, L

3Y, A

24

1971.18

406-422

A, L

L, I, Y

25

1982.01

330-345

F, L

3Y, L, A

26

1983.05

370-386

Y, L

2L, F

27

2024.02

488-505

F, L

2A, L, Y, F

28

2230.17

103-122

F, L

3L

29

2568.19

302-324

L, A

3A, L, F, Y

30

2612.41

529-550

L, F

3L, A

31

2648.38

506-528

L, L

2F, 2A, 2I, L

32

2655.4

327-348

L, L

3L, 3Y, F, A

33

2806.46

1-24

L

2F, 2L, I, A

34

2929.59

529-553

L, F

3L, 2A, F

35

2998.47

25-49

L, F

3L, 2F, A, I, Y

36

3206.59

460-487

L, F

2L

37

3269.73

330-356

F, L

4L, 4Y, 3A

38

3573.72

423-454

L, L

2L, Y

39

3616.74

226-257

A, A

4L, F, A

40

3799.88

423-456

L, L

3L, Y, I

41

3815.99

455-487

L, F

4L, I

42

4510.23

104-141

L, I

4L, 2F, 2Y, A, W

43

5211.6

460-505

L, L

3L, 2F, 2A, Y

44

5241.77

506-550

L, F

5L, 3A, 2F, 2I

45

5254.58

155-200

L, A

5A, 4L, 3Y, F, I

46

5412.79

407-454

L, L

3L, 2Y, I

47

5558.94

506-553

L, F

5L, 4A, 3F, 2I

48

5820.99

455-505

L, L

5L, 2A, 2F, Y, I

49

6124.03

103-153

F, L

5L, 4Y, 3F, 3A ,W ,I

50

6237.16

103-154

F, L

6L, 4Y, 3F, 3A, W, I

51

7174.47

346-407

L, I

8L, 5A, 3Y, 3F, I

52

9024.38

283-361

L, A

9L, 9A, 5Y, 3F, 2I

150

BSA sequence:
1

DTHKSEIAHRFKDLGEEHFKGLVLIAFSQYLQQCPFDEHVKLVNELTEFAKTCVA

DESHAGCEKSLHTLFGDELCKVASLRETYGDMADCCEKQEPERNECFLSHKDDSP
DLPKLKPDPNTLCDEFKADEKKFWGKYLYEIARRHPYFYAPELLYYANKYNGVFQ
ECCQAEDKGACLLPKIETMREKVLASSARQRLRCASIQKFGERALKAWSVARLSQ
KFPKAEFVEVTKLVTDLTKVHKECCHGDLLECADDRADLAKYICDNQDTISSKLKE
CCDKPLLEKSHCIAEVEKDAIPENLPPLTADFAEDKDVCKNYQEAKDAFLGSFLYE
YSRRHPEYAVSVLLRLAKEYEATLEECCAKDDPHACYSTVFDKLKHLVDEPQNLIK
QNCDQFEKLGEYGFQNALIVRYTRKVPQVSTPTLVEVSRSLGKVGTRCCTKPESER
MPCTEDYLSLILNRLCVLHEKTPVSEKVTKCCTESLVNRRPCFSALTPDETYVPKAF
DEKLFTFHADICTLPDTEKQIKKQTALVELLKHKPKATEEQLKTVMENFVAFVDK
CCAADDKEACFAVEGPKLVVSTQTALA

We also employed trypsin/PSS-modified membranes to digest BSA denatured with a
NALS, but there is no obvious trend in peptide size with residence time. With a ~0.05-s
digestion, the sequence coverage is 90%, even higher than for short-time peptic digestion. The
relatively high sequence coverage with the tryptic digest may stem from a denaturing procedure
that, in contrast to the denaturation for peptic digestion, included alkylation and no urea.
However, out of 65 tryptic peptides, 63 have masses less than 5 kD and the remaining 2 peptides
have masses between 5 and 6 kD. Large peptides are important for correlating neighboring
posttranslational modifications.[22-23] Unfortunately, with the simple syringe pump system we
could not achieve accurate flow control at higher rates where larger tryptic peptides might appear.
Although pepsin provides large peptides more easily than trypsin, low specificity is a
drawback of pepsin. In our database searching, we assume peptic cleavage occurs at the C151

terminus side of A, F, I, L, W and Y.

For apomyoglobin digestion in pepsin-modified

membranes, we observe at least some cleavage at 2/15 A, 1/7 F, 1/9 I, 9/17 L, 0/2 W and 0/2 Y
residues. With BSA, the values are 8/47 A, 12/27 F, 2/14 I, 28/61 L, 0/2 W and 1/20 Y residues.
In both cases pepsin cleavage occurs preferentially after L, and with BSA, peptic cleavage after
F also occurs at nearly 50% of the sites. These data are consistent with a previous statistical
analysis of the specificity of immobilized pepsin in the digestion of 39 proteins (13766 amino
acids residuals). That study shows that F and L have a cleavage probability larger than 40 %,
whereas cleavage is less than 20% at A, I, W, and Y sites. [24] One particular challenge in
working with pepsin is that other cleavage sites or N-terminus cleavage of A, L, I, F, W or Y
could lead to unassigned peptide signals, but tandem mass spectrometry can potentially
overcome this problem.[25]

4.4

Conclusions
Immobilizing common proteases (trypsin and pepsin) in nylon membranes at high
3

enzyme density (30-60 mg/cm ) conveniently creates membrane-based enzymatic reactors.
These reactors catalyze limited proteolysis in <1 s. More importantly, tuning the flow rate
through the membrane affords control over the size of proteolytic peptides. For apomyoglobin
digestion by pepsin, shorter residence times yield longer peptides with more charge. A large
peptide (8kD) with a +10 charge state provides a spatial resolution of <2 amino acid residues in
ETD MS/MS analysis. Similarly, in membrane-based peptic digestion of denatured BSA, limited
digestion yields many missed cleavages and creates long peptides that increase the sequence
coverage from 53% to 82% on going from 2-s to 0.05-s digestion. Future work will focus on

152

limited digestion of proteins with multiple posttranslational modifications to study correlations
between different modifications.

4.5

Acknowledgement

We are grateful to the U.S. National Institutes of Health (GM 080511 to M.B.) for funding this
work. I would like to thank Ms. Li Cui from Prof. Gavin Reid’s group for the initial help of using
the Orbitrap Velos instrument.

153

REFERENCES

154

4.6

References

1.

Ma, J.; Zhang, L.; Liang, Z.; Shan, Y.; Zhang, Y., Immobilized Enzyme Reactors in
Proteomics. TrAC, Trends Anal. Chem. 2011, 30 (5), 691-702.

2.

Ma, J.; Hou, C.; Sun, L.; Tao, D.; Zhang, Y.; Shan, Y.; Liang, Z.; Zhang, L.; Yang, L.;
Zhang, Y., Coupling Formic Acid Assisted Solubilization and Online Immobilized
Pepsin Digestion with Strong Cation Exchange and Microflow Reversed-Phase Liquid
Chromatography with Electrospray Ionization Tandem Mass Spectrometry for Integral
Membrane Proteome Analysis. Anal. Chem. 2010, 82 (23), 9622-9625.

3.

Rey, M.; Man, P.; Brandolin, G.; Forest, E.; Pelosi, L., Recombinant Immobilized
Rhizopuspepsin as a New Tool for Protein Digestion in Hydrogen/Deuterium Exchange
Mass Spectrometry. Rapid Commun. Mass Spectrom. 2009, 23 (21), 3431-3438.

4.

Gao, J.; Xu, J.; Locascio, L. E.; Lee, C. S., Integrated Microfluidic System Enabling
Protein Digestion, Peptide Separation, and Protein Identification. Anal. Chem. 2001, 73
(11), 2648-2655.

5.

Krenkova, J.; Lacher, N. A.; Svec, F., Highly Efficient Enzyme Reactors Containing
Trypsin and Endoproteinase LysC Immobilized on Porous Polymer Monolith Coupled to
MS Suitable for Analysis of Antibodies. Anal. Chem. 2009, 81 (5), 2004-2012.

6.

Cooper, J. W.; Chen, J.; Li, Y.; Lee, C. S., Membrane-Based Nanoscale Proteolytic
Reactor Enabling Protein Digestion, Peptide Separation, and Protein Identification Using
Mass Spectrometry. Anal. Chem. 2003, 75 (5), 1067-1074.

7.

Dulay, M. T.; Baca, Q. J.; Zare, R. N., Enhanced Proteolytic Activity of Covalently
Bound Enzymes in Photopolymerized Sol Gel. Anal. Chem. 2005, 77 (14), 4604-4610.

8.

Xu, F.; Wang, W. H.; Tan, Y. J.; Bruening, M. L., Facile Trypsin Immobilization in
Polymeric Membranes for Rapid, Efficient Protein Digestion. Anal. Chem. 2010, 82 (24),
10045-10051.

9.

Walter, B., Characterization of Agarose-Bound Trypsin. Biochim. Biophys. Acta 1976,
429 (3), 950-953.

155

10.

Liuni, P.; Rob, T.; Wilson, D. J., A Microfluidic Reactor for Rapid, Low-Pressure
Proteolysis with On-Chip Electrospray Ionization. Rapid Commun. Mass Spectrom. 2010,
24 (3), 315-320.

11.

Engen, J. R., Analysis of Protein Conformation and Dynamics by Hydrogen/Deuterium
Exchange MS. Anal. Chem. 2009, 81 (19), 7870-7875.

12.

Konermann, L.; Pan, J.; Liu, Y.-H., Hydrogen Exchange Mass Spectrometry for Studying
Protein Structure and Dynamics. Chem. Soc. Rev. 2011, 40 (3), 1224-1234.

13.

Ge, Y.; Rybakova, I. N.; Xu, Q.; Moss, R. L., Top-down High-resolution Mass
Spectrometry of Cardiac Myosin Binding Protein C Revealed that Truncation Alters
Protein Phosphorylation State. Proc. Natl. Acad. Sci. U. S. A. 2009, 106 (31), 1265812663.

14.

Liu, G.; Dotzauer, D. M.; Bruening, M. L., Ion-Exchange Membranes Prepared using
Layer-by-Layer Polyelectrolyte Deposition. J. Membr. Sci. 2010, 354 (1–2), 198-205.

15.

Bhattacharjee, S.; Dong, J.; Ma, Y.; Hovde, S.; Geiger, J. H.; Baker, G. L.; Bruening, M.
L., Formation of High-Capacity Protein-Adsorbing Membranes through Simple
Adsorption of Poly(acrylic acid)-Containing Films at Low pH. Langmuir 2012, 28 (17),
6885-6892.

16.

Bi, H.; Qiao, L.; Busnel, J.-M.; Liu, B.; Girault, H. H., Kinetics of Proteolytic Reactions
in Nanoporous Materials. J. Proteome Res. 2009, 8 (10), 4685-4692.

17.

Fontana, A.; Zambonin, M.; Polverino de Laureto, P.; De Filippis, V.; Clementi, A.;
Scaramella, E., Probing the Conformational State of Apomyoglobin by Limited
Proteolysis. J. Mol. Biol. 1997, 266 (2), 223-230.

18.

Syka, J. E. P.; Coon, J. J.; Schroeder, M. J.; Shabanowitz, J.; Hunt, D. F., Peptide and
Protein Sequence Analysis by Electron Transfer Dissociation Mass Spectrometry. Proc.
Natl. Acad. Sci. U. S. A. 2004, 101 (26), 9528-9533.

19.

Good, D. M.; Wirtala, M.; McAlister, G. C.; Coon, J. J., Performance Characteristics of
Electron Transfer Dissociation Mass Spectrometry. Mol. Cell. Proteomics 2007, 6 (11),
1942-1951.

156

20.

Pan, J.; Han, J.; Borchers, C. H.; Konermann, L., Hydrogen/Deuterium Exchange Mass
Spectrometry with Top-Down Electron Capture Dissociation for Characterizing
Structural Transitions of a 17 kDa Protein. J. Am. Chem. Soc. 2009, 131 (35), 1280112808.

21.

Wu, S.-L.; Kim, J.; Hancock, W. S.; Karger, B., Extended Range Proteomic Analysis
(ERPA):  A New and Sensitive LC−MS Platform for High Sequence Coverage of
Complex Proteins with Extensive Post-translational ModificationsComprehensive
Analysis of Beta-Casein and Epidermal Growth Factor Receptor (EGFR)†. J. Proteome
Res. 2005, 4 (4), 1155-1170.

22.

Kellie, J. F.; Tran, J. C.; Lee, J. E.; Ahlf, D. R.; Thomas, H. M.; Ntai, I.; Catherman, A.
D.; Durbin, K. R.; Zamdborg, L.; Vellaichamy, A.; Thomas, P. M.; Kelleher, N. L., The
Emerging Process of Top Down Mass Spectrometry for Protein Analysis: Biomarkers,
Protein-Therapeutics, and Achieving High Throughput. Mol. Biosyst. 2010, 6 (9), 15321539.

23.

Siuti, N.; Kelleher, N. L., Decoding Protein Modifications using Top-Down Mass
Spectrometry. Nat. Methods 2007, 4 (10), 817-821.

24.

Hamuro, Y.; Coales, S. J.; Molnar, K. S.; Tuske, S. J.; Morrow, J. A., Specificity of
Immobilized Porcine Pepsin in H/D Exchange Compatible Conditions. Rapid Commun.
Mass Spectrom. 2008, 22 (7), 1041-1046.

25.

Nelson, D. L.; Lehninger, A. L.; Cox, M. M., Lehninger Principles of Biochemistry. W.H.
Freeman: New York, 2008.

157

Chapter 5. Protease-Modified Membranes for Studies of
Protein Structure: Limited Proteolysis and Rapid Digestion
for HDX
(The work on RHD3 is part of a manuscript submitted to Analytical Chemistry)
Chapter 4 introduced protease-modified membranes whose low thickness (170 m)
enables limited digestion with msec residence times. This chapter explores the use of proteasemodified membranes as part of two techniques for investigating protein structure, HDX and
limited digestion of proteins without denaturation.

After HDX, rapid peptic digestion in

membranes facilitates MS analysis of the mass increase on deuterium-containing peptides. In
contrast, limited proteolysis of a large Arabidopsis GTPase, ROOT HAIR DEFECTIVE 3
(RHD3), shows suitable probing for labile regions near the C-terminus to suggest what protein
reconstruction might make RHD3 more suitable for crystallization.

5.1

Introduction
Complementary to NMR and X-ray crystallography, HDX followed by MS is an

emerging method in the study of protein structure and dynamics.[1] The sites of deuterium
exchange on a protein after exposure to D2O often indicate which regions of a protein or
protein complex are accessible to water and have weak hydrogen bonding.

Moreover, the

kinetics of exchange can also suggest the time-frame of protein dynamics.[2] After a specified
time of protein exposure to D2O, quenching of HDX occurs via acidification and a decrease in
temperature.

Moreover, the low pH (pH=2.5) and temperature (0 º are also important for
C)
158

preventing back-exchange during digestion and analysis.[1, 3] Due to the need for low pH,
pepsin is the ideal protease for HDX applications.[4] In this chapter, after exposure to D2O for 5
min we digest apomyoglobin in pepsin-modified membranes (1-s residence time) at pH 2.5 and
subject the resulting peptides to ESI-MS analysis (Figure 5.1). Comparison of the spectra from
digests of deuterium-exchanged and control proteins shows the mass increase due to the
replacement of amide hydrogen by deuterium at different regions. (The technique must take into
account exchange on the side chains of labile residues.)
Limited digestion can also provide low-resolution structural information when digesting
proteins in their native configuration. In this case proteolysis only occurs in target protein
regions that are readily accessible to the enzyme. Traditional limited protein digestion employs a
solution-based approach, which includes varying the incubation time, enzyme to substrate ratio,
or reaction temperature to control the extent of proteolysis. This approach is laborious and
sometimes difficult to reproduce.[5-6] With protease-containing membranes, similar experiments
can be performed by simply varying the flow rate of a protein solution through the membrane.
This method can greatly decrease the experimental time (~15 min) and sample consumption
(~100 g) in limited digestion studies, and fine control of the flow rate using a syringe pump
may increase reproducibility. We demonstrate this concept by investigating which regions of
recombinant RHD3 are flexible and susceptible to hydrolysis. Based on sodium dodecyl sulfate
polyacrylamide gel electrophoresis (SDS-PAGE) and MS data, we find four readily accessible
regions at the protein C-terminus and suggest that truncation of the most labile site may increase
the likelihood of RHD3 crystallization.

159

5 mM NH4HCO3 pH=7.8,
90%D2O, 5 min HDX

Quench with formic
acid to pH 2.5.
Proteins undergo
membrane digestion
in a 1-s residence time.

5 mM NH4HCO3 pH=7.8,
H2O, 5 min control
0.1 mg/mL apomyoglobin

Figure 5.1. The workflow of HDX and membrane digestion. The 0.1 mg/mL apomyoglobin is
exchanged in 90% D2O (or 100% H2O) for 5 min (5 mM NH4HCO3, pH=7.8). The reaction is
quenched by lowering the pH to 2.5 via addition of FA, and the deuterium protein or control
protein (no exchange) immediately undergoes membrane digestion to generate peptides with or
without deuterium.

5.2

Experimental

5.2.1 Materials, membranes and membrane holders
See chapter 4 for details in this area.

5.2.2 Membrane modification with protease
The modification of membranes with pepsin and trypsin was the same as described in
chapter 4. An -chymotrypsin-modified membrane was also used in the RHD3 study. For chymotrypsin immobilization, ten mL of a 0.02 M PSS, 0.5 M NaCl solution (pH=2.3) was first
passed through the membrane at 2 mL/min, followed by 30 mL of deionized water at the same
flow rate. Subsequently, 5 mL of 1 mg/mL -chymotrypsin in 2.7 mM HCl, 2 mM CaCl2 was
circulated through the membrane at 1 mL/min for one hour. After protease deposition, the
membrane was rinsed with 30 mL of 1 mM HCl. Modified membranes were dried and stored in a
160

desiccator. The amounts of -chymotrypsin immobilized in membranes were determined from
direct UV absorbance at 280 nm using a Nanodrop

TM

UV/Vis Spectrometer (Thermo).

5.2.3 Apomyoglobin HDX and membrane-digestion procedure
One mg of apomyoglobin was dissolved in one mL of 50 mM NH4CO3 to make a stock
solution. One hundred L of stock solution was diluted with D2O or H2O (control) at a 1:9 ratio,
and the mixture was left for 5 min at room temperature. The reaction was quenched by addition
of 50 L of formic acid (FA), and 100 L of the quenched sample (0.095 mg/mL or 6 M
apomyoglobin) was immediately passed through the pepsin-modified membrane at a flow rate of
0.6 mL/h (1 s residence time), and this process took 10 min. The 100 L of digested sample was
mixed with 100 L acetonitrile and frozen quickly with liquid nitrogen prior to lyophilization
with a Speed-Vac. The lyophilization took 20 min, during which the mixture was frozen. The
dried digest was dissolved in 100 L of MS buffer (3% FA, 47% H2O and 50% methanol,
pH=2.5, precooled on ice), and 40 L of this sample was infused into ESI-MS. The time from
dissolving to injection was 3 min.

5.2.4 RHD3 expression and purification
The RHD3(1-676)-HIS6 was cloned by Dr. Giovanni Stefano from Prof. Federica
Brandizzi’s group, and the protein expression and purification was performed by Dr. Yi Zheng
from Prof. R. Michael Garavito’s group. The Production of RHD3(1-676)-HIS6 subcloned in
pET28b vector was induced in Escherichia coli (BL21 λDE3). The bacteria were cultured at
37 ° in LB medium containing 100 g/ml of kanamycin until reaching an optical density of 1.0
C
at 600 nm.

Protein production was induced by addition of 1 mM isopropyl 1-thio-β-D161

galactopyranoside for 5 h at 30 ° Harvested cells from 1 L of cell culture were suspended in 50
C.
mL of buffer A (30 mM Tris-HCl, pH 8.0, 200 mM NaCl, 4 mM MgCl2, 2 mM βmercaptoethanol and 1 mM phenylmethanesulfonyl fluoride ) and lysed with 4 min of sonication
at an output of 48 watts with 10 s pulses (Sonicator Misonix 3000). Following a centrifugation
step at 20,000 x g (4 ° for 20 min, the clarified lysate was applied to a 7-mL Ni-NTA (Qiagen)
C)
gravity flow column, which was pre-equilibrated in buffer A containing 25 mM imidazole. After
extensive washing with buffer A, fractions were eluted over 2 column volumes using buffer A
containing 150 mM imidazole. Fractions containing RHD3 (as shown by UV absorbance at 280
nm) were pooled and concentrated to about 1 mL, and 500 µL volumes were applied at 0.4
ml/min to a Superose 6 10/300 GE gel filtration column (GE Healthcare), which was preequilibrated with buffer B (30 mM Tris-HCl, pH 8.0, 200 mM NaCl, 4 mM MgCl2, 1 mM
EDTA, and 1 mM DTT). Peak fractions containing target proteins were concentrated using a
spin column (30 kD molecular weight cutoff) to ~15 mg/mL based on UV/Vis absorbance at 280
nm. Analytical gel filtration chromatography was also used to estimate the molecular size of
purified RHD3, which eluted at a position corresponding to a homo dimer with a molecular
weight of 150 kD.

5.2.5 Limited proteolysis of RHD 3 with either trypsin- or -chymotrypsinmodified membranes
The gel filtration-purified RHD3 was diluted with 10 mM NH4HCO3 to make 1 mL of
0.1 mg/mL RHD3, and this solution was passed through a small trypsin- or -chymotrypsinmodified membrane at 0.6, 3 and 13 mL/h. The digests were analyzed by MALDI-MS and ESI-

162

MS, and 3 g of each digested protein and intact protein were also analyzed by SDS-PAGE with
Coomassie Blue staining.

5.2.6 Mass spectrometry and data analysis
See chapter 4 for details in this area.

5.3

Results and discussion

5.3.1 Preliminary study of HDX on apomyoglobin using digestion in a pepsinmodified membrane
In HDX, digestion of the deuterium-exchanged protein should be rapid to prevent back
exchange in H2O matrix, or in our case to prevent over-exchange in a D2O matrix. Highly
active pepsin-containing reactors can catalyze the digestion in a short time. Moreover the
immobilized-pepsin reactor keeps the enzyme away from the peptic digest after proteolysis to
decrease MS interferences from self-digested enzyme.[1, 4] Pepsin-modified membranes may be
especially effective reactors to digest deuterium-exchanged proteins because of the short radial
diffusion distances in membrane pores. We performed membrane digestion of deuteriumexchanged apomyoglobin using 1-s residence times and analyzed the peptic digests using ESIMS. Figures 5.2a and 5.2b show the spectra of 6 M peptic digests of deuterium-exchanged
apomyoglobin and control (non-exchanged) apomyoglobin, respectively. They demonstrate very
similar digestion, so it is easy to compare the two spectra. As expected, the isotopic distributions
of peptide signals in 5.2a are wider than in 5.2b, and dominant signals shift to a slightly higher
m/z value. The deuterium exchange not only increases a peptide’s mass but variable exchange
also complicates the isotopic mass distribution. Figures 5.2c and d are expanded (x-axis) views
163

of signals from a peptide selected (red rectangle) from a and b, respectively. The peptides have
the same sequence (amino acids 1-29 of apomyoglobin) and both show a monoisotopic mass at
m/z 1045.53 (+3 charge state). We use the difference between the centroid masses (shown in
Figure 5.2d) to estimate the amount of exchanged deuterium. (The centroid mass is calculated as
the peak height-weighted averaged mass of all isotopic peaks.) The 2.24 Da difference between
the centroid masses indicts an average 6.72 Da mass increase (based on the +3 charge state) due
to D exchange. The mass increases occur to different extent with different peptides. The above
data suggest the success of HDX and the peptic membrane digestion, but localization of
deuterium exchange to more specific amino acid regions needs further investigation. This will
require either ETD fragmentation or digestion into smaller, overlapping peptides.

164

a

b

650

700

750

800

850

900
m/z

950

1000 1050 1100 1150

c

d

1048.38-1046.14=2.24 Da (centroid mass difference)

1046

1047

1048

m/z

1049

1050

1051

Figure 5.2. ESI-Orbitrap mass spectra of 6 M (a) deuterium-exchanged or (b) control
apomyoglobin digested by a pepsin-modified membrane (1-s residence time). (c) and (d) are
165

enlarged (x-axis) views of the selected peaks (red rectangles) from (a) and (b), respectively. The
centroid masses of (c) and (d) are labeled by the dash line, and the difference is given.

5.3.2 Limited membrane proteolysis of RHD3 for structural information
In a simpler method for investigating protein structure with low resolution, we suggest
which regions of a protein are highly accessible to enzymes by observing where limited digestion
occurs. RHD3 serves as our initial example. Homologous to human atlastins, RHD3 is one of
the largest membrane-bound GTPases in Arabidopsis thaliana.[7]

This protein participates in

the modeling of the endoplasmic reticulum and contains a GTPase domain, a helix bundle, two
trans-membrane helices and one cytosolic tail.[8] Human atlastin may be involved in homotypic
fusion of the membrane of the endoplasmic reticulum, and much of the information in the
mechanisms that could underpin such a fusion has been gathered from crystallographic analyses
of the soluble domain (cytosolic) of atlastin.[7] The biological role of RHD3 is unknown,[9]
and a crystal structure of this protein would most likely offer valuable insights into RHD3’s role
in plant cells.
This work aims to use limited membrane-based proteolysis to map flexible protein
regions that may restrict crystallization (Scheme 5.1).[10-11] Flexible regions should contain
digestion sites that are much more accessible to proteolysis than similar sites in the globular
region of the protein. Subsequently, proteins expressed without the flexible regions would be
more stable and compact, and consequently have a much higher probability of crystallization.[56, 10, 12] Because the globular regions of the protein resist digestion, we could employ trypsin
and -chymotrypsin in limited proteolysis. These enzymes are more specific than pepsin and
operate at a non-denaturing pH (NH4HCO3 buffer, pH 7.5), which is vital for structural studies.

166

Scheme 5.1. Strategy for limited RHD3 digestion in membranes containing trypsin and chymotrypsin. The protein picture (top left) is that for a human atlastin dimer (made from PDB
3QOF and 3QNU using Pymol), which has a similar domain configuration to RHD3 but with
less -helix. The drawings represent RHD3 dimers (one GTPase domain and 9 -helices on
each monomer).
To obtain different digestion patterns, we selectively pass the RHD3 solutions through
either trypsin- or -chymotrypsin-modified membranes at 0.6, 3 and 13 mL/h. Figure 5.3 shows
the SDS-PAGE analysis of the proteolytic digests. In both images, lane 1 is the protein ladder,
lane 2 is the intact protein, and lanes 3-5 are the digests at the different flow rates, which
correspond to residence times of 0.045, 0.2 and 1 s, respectively. After tryptic digestion, three
major protein bands appear with molecular masses around 70, 65, and 60 kD. We performed the
-chymotrypsin membrane digestion one week after the trypsin membrane digestion, and the
protein itself degraded slightly, presumably because of trace protease not removed during the
167

purification process. The SDS-PAGE after chymotrypsin digestion reveals a dominant
intermediate protein with a molecular weight around 65 kD.

Figure 5.3. SDS-PAGE analysis of limited RHD3 digestions in a trypsin-modified membrane
(left) and an -chymotrypsin-modified membrane (right). Three g of either protein or digests
were loaded onto each column, and Lanes 1-5 are protein ladder, intact protein, digests at 13
mL/h (residence time of 0.045 s), digests at 3 mL/h (residence time of 0.2 s) and digests at 0.6
mL/h (residence time of 1 s), respectively.
We applied both MALDI-LTQ MS and ESI-Orbitrap MS to investigate the most
abundant peptides in the RHD3 digests. Figures 5.4a, b and c show the MALDI mass spectra of
RHD3 digested in a trypsin-modified membrane for different residence times. Peak labels show
peptide sequences determined from comparison of masses with theoretical masses obtained from
168

in silico digestion. The recombinant RHD3 protein has 691 amino acids including its
polyhistidine tag, and the most intense peak at the shortest digestion time corresponds to the
peptide with amino acids 673-691, suggesting a flexible polyhistidine tag region. The mass
spectra also indicate cleavage after amino acids R542, R560, R569, K600, K617, K632, K637,
K646 and R650. -Chymotrypsin also shows cleavage sites in these regions (Table 5.1). In the
predicted secondary structure, all of these peptides lie within the last 4 -helices or regions
between them.

Comparison of the three spectra in Figure 5.4 suggests that cleavage is most

rapid at R672, as relative to the peptide 673-691, signals for other peptides typically increase
with increasing digestion time. This suggests that longer digestion times result in digestion
deeper into the protein, which is consistent with the SDS-PAGE results.
The ESI-Orbitrap mass spectrum (Figure 5.4d) confirms MALDI-MS results and shows
dominant cleavage sites after R576, K617, R650, and R672. The signal for peptide 673-691 is
especially strong. Overall, the mass spectra suggest there are 4 highly accessible regions around
R576, K617, R650, and R672. Because ESI gives multiple charge states, we see larger peptides
than with MALDI. Figure 5.5 (ESI mass spectra of RHD3 digested in a trypsin-modified
membrane for different residence times) shows that peptide 618-691 is present primarily at low
digestion times, suggesting that the K617 site is more accessible than R650. This Figure also
reveals a small signal for peptide 1-71 from the N-terminus, showing limited access to this part
of the protein. Table 5.1 lists the most abundant peptides in limited trypsin and -chymotrypsin
digests. In Figures 5.4a, b and c there is a signal that seems to correspond to peptide 251-263.
This peptide is in the globular region of the protein where the GTPase domain and helical region
connect, and this linker region may be also accessible to the enzyme.

169

a

b

673-691

2000

600

2500
m/z

673-691
618-691
618-646
618-691
651-672
577-617
618-650
618-691

651-691

673-691

651-672

d

1500

700

800

900
m/z

3000

3500

618-646
618-672
618-691
1-71
577-617

1000

618-646

1-71

618-632
601-617
251-263
618-637 672-691
561-569
651-672
543-560

618-691

c

1000 1100 1200

Figure 5.4. MALDI mass spectra of 0.1 mg/mL (1.3 M) RHD 3 digested in residence times of
(a) 1 sec, (b) 0.2 sec and (c) 0.045 s by a trypsin-modified membrane, and (d) ESI mass spectrum
of the same sample as (c). Signals were assigned to peptides by mass fingerprinting and are
labeled with the amino acid position of these peptides (protein full length 1-691). Compared to
the C-terminus peptide (673-691, stars), the relative intensities of the smaller peptides cleaved
170

after amino acids K600, K617 (red circles), R650 (green circle) or R560 (blue circles) increase
as the residence time increases from (c) to (b) to (a), suggesting that longer digestion times result
in deeper access into the protein.

171

Figure 5.5. ESI-Orbitrap mass spectra of 0.1 mg/mL (1.3 M) RHD 3 digested in residence times
of (a) 1, (b) 0.2 and (c) 0.045 s by a trypsin-modified membrane. The peptides are labeled with
172

their amino acid positions. The dominant peak at m/z 570.78 suggests that the most labile
cleavage site is after K672, and the presence of other major peptides in (c) cleaved after amino
acids R650, K617 and R576 is consistent with the MALDI mass spectra in Figure 7a, b and c.
The y axis is limited to 30% of the signal at m/z 570.78.

Table 5.1. RHD3 peptides identified from MALDI and ESI-Orbitrap mass spectra obtained after
digestion in trypsin-modified and -chymotrypsin-modified membranes.

Peptide
Monoisotopic
Mass
2280.09
2436.19
2584.28
4845.35
1624.81
2149.10
3132.65
3674.95
6240.20
8501.28
1752.91
4406.35
1103.58
2140.99
1278.68
7680.71

Trypsin-modified membrane digestion
Peptide sequence
Amino
Acids
(R)NNNWVDKLAAALEHHHHHH(-)
(R)RNNNWVDKLAAALEHHHHHH(-)
(R)QFKNETEYTVTQAISAQEANRR(N)
(R)QFKNETEYTVTQAISAQEANRRNN
NWVDKLAAALEHHHHHH(-)
(K)SISTIDSLASSTWEK(V)
(K)SISTIDSLASSTWEKVAPEK(T)
(K)SISTIDSLASSTWEKVAPEKTLITPV
QCK(S)
(K)SISTIDSLASSTWEKVAPEKTLITPV
QCKSLWR(Q)
(K)SISTIDSLASSTWEKVAPEKTLITPV
QCKSLWRQFKNETEYTVTQAISAQE
ANRR(N)
(K)SISTIDSLASSTWEKVAPEKTLITPV
QCKSLWRQFKNETEYTVTQAISAQE
ANRRNNNWVDKLAAALEHHHHHH(-)
(K)TLTLALFNSTGNNATSK(S)
(R)SASLKLLSVMAVIRLDDELDNIEKT
LTLALFNSTGNNATSK(S)
(R)VWTGKEDIR(A)
(R)MKDRFATIFSHDSDSMPR(V)
(R)GVVPANAFAFSAK(Q)
MDAACSTQLIDGDGVFNVSGVDHFIK
EVKLDECGLSYAVVSIMGPQSSGKST
LLNHLFGTNFREMDAFRGR(S)
173

MS
Instrument

673-691
672-691
651-672
651-691

MALDI/ESI
MALDI/ESI
MALDI/ESI
ESI

618-632
618-637
618-646

MALDI/ESI
MALDI/ESI
MALDI/ESI

618-650

ESI

618-672

ESI

618-691

ESI

601-617
577-617

MALDI/ESI
ESI

561-569
543-560
252-263
1-71

MALDI
MALDI/ESI
MALDI
ESI

Table 5.1 (cont’d)
Peptide
Monoisotopic
Mass
3805.89
1214.58
2170.19
2342.11
1362.74
1316.57
1284.27
2582.24

-chymotrypsin-modified membrane digestion
Peptide sequence
Amino
Acids
(Y)TVTQAISAQEANRRNNNWVDKLAA
ALEHHHHHH(-)
(W)RQFKNETEY(T)
(W)EKVAPEKTLITPVQCKSLW(R)
(F)NSTGNNATSKSISTIDSLASSTW(E)
(W)TGKEDIRAITKM(A)
(F)SHDSDSMPRVW(T)
(M)RMKDRFATIF(S)
(M)RMKDRFATIFSHDSDSMPRVW(T)

MS
Instrument

659-691

MALDI/ESI

650-658
631-649
608-630
563-574
552-562
542-551
542-562

MALDI/ESI
MALDI/ESI
MALDI/ESI
MALDI/ESI
MALDI/ESI
MALDI/ESI
MALDI/ESI

RHD3 sequence:
1

MDAACSTQLIDGDGVFNVSGVDHFIKEVKLDECGLSYAVVSIMGPQSSGKSTLLNH
LFGTNFREMDAFRGRSQTTKGIWIARCAGIEPCTVVMDLEGTDGRERGEDDTAFE
KQSALFALAVSDIVLINMWCHDIGREQAANKPLLKTVFQVMMRLFSPRKTTLMFVI
RDKTRTPLENLEPVLREDIQKIWDSVPKPQAHKETPLSDFFNVEVVALSSYEEKEEQ
FKEQVYNLRQRFFQSVAPGGLAGDRRGVVPANAFAFSAKQMWQVIKDNKDLDLPA
HKVMVATVRCEEIANEKFSSFIANENWRELEEAVQSGPVSGFGRKLSSILQASLSEY
DTEATYFEESVRSSKRQQLQEKLLQLVQPTFQDVLGHLRAGALENFKNAFEKALD
AGEGFSSSAKSCAQSCISKFDKGCEEAVIEQAKWDTSKTREKLERDIEAHISSVRTAK
LAELTTLYESKLNVALSGPVEALLDGANDETWPAIRKLLRREGELAVYGLSNALSGF
EMDEETRSKMLADLENYARGIVETKAKEEAGRAMMRMKDRFATIFSHDSDSMPRV
WTGKEDIRAITKMARSASLKLLSVMAVIRLDDELDNIEKTLTLALFNSTGNNATSKSI
STIDSLASSTWEKVAPEKTLITPVQCKSLWRQFKNETEYTVTQAISAQEANRRNNN
WVDKLAAALEHHHHHH

Overall, these data suggest that the region around R672 is highly flexible and that
truncating the protein at R672 may increase the probability of crystallization. (Notably, we do
not detect the cleavage at K679). In fact a new construct up to R672 shows no degradation over
several weeks, but we have not yet successfully crystallized this protein. Compared to insolution limited digestion methods to suggest truncated protein sequences, the membrane method
is rapid and convenient and provides finer control over the digestion time.
174

5.4

Conclusions
The pepsin-modified membrane successfully generates peptides from deuterium-

exchanged apomyoglobin, and ESI-MS detects the mass increases for peptic digests. However,
more experiments needs to be done to improve resolution. The localization of deuterium
exchange could be achieved by ETD-MS/MS analysis of large peptides,[13] the overlapping of
small peptides from longer digest times,[4] or a combination of the two methods. Probing the
structure of BSA using HDX-MS coupled to membrane digestion might be an interesting and
challenging future work, and the comparison of the obtained HDX data with simulation data or
data from other methods would increase the impact of this work in the area of studying large
proteins with HDX.
Limited, membrane-based proteolysis shows which regions of a protein are readily
accessible to enzymes. For RHD3, digestion in a membrane-based enzymatic reactor reveals
rapid digestion at K672 and suggests that truncation of the protein at this amino acid will
stabilize RHD3. Future studies with other proteins will further demonstrate the value of limited
digestion in protease-modified membranes.

5.5

Acknowledgement

Many thanks to Dr. Giovanni Stefano, Prof. Federica Brandizzi (F.B.), Dr. Yi Zheng and Prof. R.
Michael Garavito (R.M.G) for their collaboration in the RHD3 study. We are grateful to the U.S.
National Institutes of Health (GM 080511, Bruening), the Chemical Sciences, Geosciences and
Biosciences Division, Office of Basic Energy Sciences, Office of Science, U.S. Department of
Energy (DE-FG02-91ER20021, F.B.), the National Science Foundation (MCB 0841594, F.B),
and the MSU-REF Center Program (REF03-016, R.M.G) for funding portions of this work.
175

REFERENCES

176

5.6

References

1.

Engen, J. R., Analysis of Protein Conformation and Dynamics by Hydrogen/Deuterium
Exchange MS. Anal. Chem. 2009, 81 (19), 7870-7875.

2.

Konermann, L.; Pan, J.; Liu, Y.-H., Hydrogen Exchange Mass Spectrometry for Studying
Protein Structure and Dynamics. Chem. Soc. Rev. 2011, 40 (3), 1224-1234.

3.

Morgan, C. R.; Engen, J. R., Investigating Solution-Phase Protein Structure and
Dynamics by Hydrogen Exchange Mass Spectrometry. In Current Protocols in Protein
Science, John Wiley & Sons, Inc.: 2009.

4.

Rey, M.; Man, P.; Brandolin, G.; Forest, E.; Pelosi, L., Recombinant Immobilized
Rhizopuspepsin as a New Tool for Protein Digestion in Hydrogen/Deuterium Exchange
Mass Spectrometry. Rapid Commun. Mass Spectrom. 2009, 23 (21), 3431-3438.

5.

Gao, X.; Bain, K.; Bonanno, J.; Buchanan, M.; Henderson, D.; Lorimer, D.; Marsh, C.;
Reynes, J.; Sauder, J.; Schwinn, K.; Thai, C.; Burley, S., High-Throughput Limited
Proteolysis/Mass Spectrometry for Protein Domain Elucidation. J. Struct. Funct.
Genomics. 2005, 6 (2), 129-134.

6.

Dong, A.; Xu, X.; Edwards, A. M., In Situ Proteolysis for Protein Crystallization and
Structure Determination. Nat. Methods 2007, 4 (12), 1019-1021.

7.

Bian, X.; Klemm, R. W.; Liu, T. Y.; Zhang, M.; Sun, S.; Sui, X.; Liu, X.; Rapoport, T. A.;
Hu, J., Structures of the Atlastin Gtpase Provide Insight into Homotypic Fusion of
Endoplasmic Reticulum Membranes. Proc. Natl. Acad. Sci. U. S. A. 2011, 108 (10),
3976-3981.

8.

Chen, J.; Stefano, G.; Brandizzi, F.; Zheng, H., Arabidopsis RHD3 Mediates the
Generation of the Tubular ER Network and Is Required for Golgi Distribution and
Motility in Plant Cells. J. Cell Sci. 2011.

9.

Hu, J.; Prinz, William A.; Rapoport, Tom A., Weaving the Web of ER Tubules. Cell
2011, 147 (6), 1226-1231.

177

10.

Cohen, S. L.; Chait, B. T.; Ferré
-D'Amaré A. R.; Burley, S. K., Probing the Solution
,
Structure of the DNA-Binding Protein Max by a Combination of Proteolysis and Mass
Spectrometry. Protein Sci. 1995, 4 (6), 1088-1099.

11.

Cohen, S. L.; Chait, B. T., Mass Spectrometry as a Tool for Protein Crystallography.
Annu. Rev. Biophys. Biomol. Struct. 2001, 30 (1), 67-85.

12.

Zhang, G.; Liu, Y.; Qin, J.; Vo, B.; Tang, W.-J.; Ruoho, A. E.; Hurley, J. H.,
Characterization and Crystallization of a Minimal Catalytic Core Domain from
Mammalian Type II Adenylyl Cyclase. Protein Sci. 1997, 6 (4), 903-908.

13.

Pan, J.; Han, J.; Borchers, C. H.; Konermann, L., Hydrogen/Deuterium Exchange Mass
Spectrometry with Top-Down Electron Capture Dissociation for Characterizing
Structural Transitions of a 17 Kda Protein. J. Am. Chem. Soc. 2009, 131 (35), 1280112808.

178

Chapter 6. Summary and Future Work
6.1

Research summary
This dissertation describes the development of functional nylon membranes containing

TiO2 nanoparticles or proteases and their applications in phosphopeptide enrichment and limited
protein digestion prior to MS analysis, respectively. Chapters 2 and 3 describe research with
TiO2-modified nylon membranes in phosphopeptide enrichment, and chapters 4 and 5 discuss
applications of protease-containing membranes in limited digestion.
Chapter 2 focuses on the characterization of TiO2-modified nylon membranes and
enrichment of model phosphopeptides and digested phosphoproteins. Alternating adsorption of
PSS and TiO2 allows rapid modification of nylon membranes. The size of aggregated TiO2
particles is about 50 nm as measured by TEM, and SEM-EDX analyses show that TiO2 adsorbs
uniformly across the membrane. Based on phosphoangiotensin breakthrough curves during
membrane loading, the TiO2-modified membranes have a binding capacity of 540 nmols of
phosphopeptide in a 22-mm diameter disc. These membranes selectively isolate the
phosphopeptides from -casein, -casein and ovalbumin digests, and even in a 100-fold excess
of non-phosphorylated BSA digests, the membrane still captures phosphopeptides from the three
model phosphoproteins digests when using 2% TFA in 1:1 H2O/acetonitrile as a loading and
washing buffer. The selectivity of the TiO2-modified membrane is sufficient for capturing the
phosphopeptides from modestly complex mixtures.

179

Chapter 3 describes the use of the TiO2-modified membranes in phosphopeptide
enrichment from immunoprecipitates and in vivo phosphorylated proteins including fusion
proteins. Enrichment with the modified membranes is vital for identifying phosphopeptides from
a digest of a tau protein, which is in vivo phosphorylated by glycogen synthase kinase-3GSK3The MALDI-MS and ESI-MS analyses of enriched p-tau phosphopeptides reveal seven
phosphorylation sites, three of which seem dominant. Comprehensive CID-, HCD- and ETDMS/MS studies localize and confirm the seven sites. In analyses of phosphopeptides from an
immunoprecipitate of p65 (from the nuclear extract of human acute monocytic leukemia cells
stimulated with tumor necrosis factor), membrane enrichment combined with MALDI-MS/MS
reveals eight phosphopeptides from the p65-associated 60S acidic ribosomal protein.
Unfortunately, none of the phosphopeptides originate from p65 and the eight peptides contain
only two unique phosphorylation sites because the peptides share a similar sequence with
different modifications from the alkylation process. In a complementary study, HPLC-ESIMS/MS analysis confirms the phosphorylation of Ser 32 and 36 on the inhibitor of kappa-Balpha protein (IB), which is in vitro phosphorylated by G-protein coupled receptor kinase-5,
and the ratio of phosphopeptide signals to their non-phosphorylated counterparts suggests ~40%
efficiency for this phosphorylation event. However, the membrane enrichment pretreatment did
not reveal any phosphorylation sites, presumably because the matrix interfered with adsorption.
Although these studies show the effectiveness of membrane enrichment, both the protein
composition and the sample matrix are important issues. For example, digestion of p65 may
only give large phosphopeptides that are not detectable in our MALDI-MS instrument.
Moreover, the large excess of ATP in studies of IB may prevent capture of phosphopeptides,

180

and detergent could also interfere with phosphopeptide adsorption. Future studies of membrane
stability will also be important for potential online applications.
Chapter 4 applies a similar adsorption procedure for creating protease-modified
membranes for protein digestion. Based on the work of Dr. Fei Xu and Dr. Wei-Han Wang with
trypsin-containing membranes, I expand the types of proteases adsorbed in membranes. The
protease-modified membranes have an enzyme density of 30 mg/cm3 (trypsin and 3

chymotrypsin) and 60 mg/cm (pepsin), and this enables the membrane to catalyze digestion
even in <1 s. The digestion occurs in membrane pores via simply passing the protein solution
through the membrane with a syringe pump. Most importantly in this work, the proteasecontaining membrane affords control of peptides sizes by altering the flow rate.

Even though

digestion is rapid, because the membranes are only 170 m thick, the residence time can be as
low as a few msec to limit the extent of digestion. For digestion with pepsin-modified
membranes, ESI-MS analysis demonstrates that the sizes of peptic peptides increase as the
residence time decreases for both BSA and apomyoglobin. In the case of BSA membrane
digestion, the sequence coverage increases from 53% (2-s digestion) to 82% (0.05-s digestion).
Additionally the ETD-MS/MS analysis of a large apomyoglobin peptide (8 kD) provides a 1-2
amino acid resolution, which suggests that this protease-modified membrane might find its
application in hydrogen-deuterium exchange (HDX) or middle-down proteomics studies.
Chapter 5 explores applications of the protease-modified membranes for probing protein
structure, both from the perspective of limited proteolysis to reveal accessible protein regions or
rapid peptic digestion for HDX studies.

Using trypsin- and -chymotrypsin-modified

membranes, the limited proteolysis of a large Arabidopsis GTPase, ROOT HAIR DEFECTIVE 3
(RHD3), shows suitable probing for labile regions near the C-terminus to suggest the protein
181

reconstruction. Truncation of a protein construct at the most labile digestion site, K672, gives a
protein that is stable in solution for a few weeks without observable degradation. This, along
with the absence of a flexible region should increase the likelihood of the crystallization of this
target protein.
Preliminary ESI-MS analysis of digests of deuterium-exchanged apomyoglobin suggests
the success of deuterium exchange on apomyoglobin and the following digestion on the pepsinmodified membrane. However we have not yet investigated localization of the deuterium
exchange, and we need to validate whether there is over-exchange or back-exchange occurring
during the workflow. There are two options to increase the resolution, and one is based on the
ETD-MS/MS analysis of the large peptides from limited digestion. However, some portions of
the sequence are only represented by small peptides that are not amenable to ETD. Another
option to enhance amino acid resolution is to increase the digestion efficiency and generate many
small peptides with overlapping sequences. The overlapped deuterium exchange states from MS
can help to increase the resolution. Slower digestion in membranes combined with multiple
enzymes might successfully yield the small overlapping peptides. Additionally, limited digestion
and ETD may be more attractive for large (>50 kD) proteins.

6.2

Future work

6.2.1 Study of the correlation of multiple phosphorylation events on p-tau
and -synuclein using membrane enrichment combined with limited
digestion or intact protein analysis
The investigation of p-tau phosphorylation in chapter 3 reveals seven phosphorylation
sites after enrichment, and three sites (S396, S400 and S404) seem to dominate the GSK182

3phosphorylation events in our study. There are 23 phosphorylation sites reported in various
studies of p-tau phosphorylated by GSK-3[1]  The full phosphorylation mapping may prove
important in understanding p-tau behavior in Alzheimer’s disease. Nevertheless, the correlation
between the phosphorylation of tau and its aggregation into paired helical filaments and
neurofibrillary tangles is still hard to assess. Mutation of the vital site which triggers or
accelerates the aggregation process may abolish or alleviate the aggregation. However among all
the phosphorylation sites, only a few may play vital roles in aggregation. We suspect that the
sites with higher phosphorylation stoichiometry are more likely to serve as the key factors in
aggregation. Thus proving that the three phosphorylation sites (S396, S400 and S404) are indeed
those with the highest stoichiometry may have relevance to understanding p-tau aggregation.
Measurement of the exact mass of intact p-tau can potentially demonstrate the average
number of phosphorylation sites per protein. Presuming this number is 3, it would be consistent
with S396, S400 and S404 being the dominant sites of phosphorylation. However, the isotopic
resolution for intact protein (>50 kD) is still challenging for our current ESI-Orbitrap instrument,
especially when the protein is not homogenous (various modifications). Adequate starting
material, careful sample desalting and optimization of mass spectrometer settings are
prerequisites to get an accurate mass increase due to phosphorylation. In a complementary
method based on the strategy of middle-down proteomics, a large peptide covering multiple
posttranslational modification (PTMs) sites will provide the correlation information for these
PTMs.[2-3] For this particular study, the peptide

386

TDHGAEIVYKpSPVVpSGDTpSPR and

407

HLpSNVSSTGSIDMVDpSPQLATLADESASLAK happen to be successive in the p-tau and

separated due to the cleavage at R406. Assuming that the limited proteolysis on the trypsin
membrane

would

possibly

miss

the

R406
183

site,

we

expect

to

observe

less

386

TDHGAEIVYKpSPVVpSGDTpSPRHLpSNVSSTGSIDMVDpSPQLATLADESASLAK(5

P)

and

more

386

TDHGAEIVYKpSPVVpSGDTpSPRHLSNVSSTGSIDMVDSPQLATLADESASLAK(3P)

if the S396, S400 and S404 are major phosphorylation sites while the S409 and S422 are minor.
Preliminary data suggest the possibility of limited digestion for looking at the abundance
of phosphorylation events. Figure 6.1 shows the MALDI mass spectra of 2 pmol of p-tau
digested for (a) 0.05 s in a trypsin-modified membrane and (b) 16 h in solution. The spectra
show some similar peptides, but new peptides also emerge from the limited digestion, suggesting
missed cleavages. Unfortunately, the mass range does not show peptides large than 4 kD. The
use of enrichment combined with ESI-MS will reveal more information from the limited digests
of p-tau. Besides p-tau, -synuclein, another important protein associated with neurological
diseases is also available from our collaborator Prof. Min-Hao Kuo. This protein has a molecular
weight of 14 kD and three potential phosphorylation sites (Figure 6.2). Future work will focus on
correlating phosphorylation events on p-tau and -synuclein using intact protein analysis and
limited digestion with phosphopeptide enrichment.

184

a

b

1000

1500

2000

2500
m/z

3000

3500

4000

Figure 6.1. MALDI mass spectra of 2 pmol of p-tau digested for (a) 0.05 s in a trypsin-modified
membrane and (b) 16 h in solution.

Alpha-Synuclein, associated with Parkinson's disease

Size: 140 Amino Acids MW: 14460 Da
Potential phosphorylation sites: S87, Y125, S129
10
20
30
40
50
60
MDVFMKGLSK AKEGVVAAAE KTKQGVAEAA GKTKEGVLYV GSKTKEGVVH GVATVAEKTK
70
80
90
100
110
120
EQVTNVGGAV VTGVTAVAQK TVEGAGSIAA ATGFVKKDQL GKNEEGAPQE GILEDMPVDP
130
140
DNEAYEMPSE EGYQDYEPEA

Figure 6.2. Properties of -synuclein.
185

6.2.2 Phosphorylation mapping in tau isomers and multiple kinase events
To date, multiple approaches such as MS, western blotting, and Edman degradation have
revealed more than 40 phosphorylation sites in tau protein associated with Alzheimer’s disease,
resulting from in vitro phosphorylation of tau isomers by four kinases.[1] The four pathological
kinases in humans are GSK-3, cyclin-dependent kinase-5, casein kinase 1 and cyclic adenosine
monophosphate-dependent protein kinase, and tau isomers in humans are differentiated into six
(0N3R, 0N4R, 1N3R, 1N4R, 2N3R, 2N4R) variations based on the presence or absence
of specific exons at the C-terminus or N-terminus.[4] To examine phosphorylation in in vivo
studies, our collaborator Prof. Kuo employs the protein interaction module-assisted catalysis
technique (Figure 6.3).

In chapter 3 we studied p-tau (1N4R) phosphorylation by GSK-

3however the enzyme and substrate can be replaced by any of the other candidates to reveal
other phosphorylation sites. In addition to the enzyme/substrate replacement, the affinity zipper
Jun-Fos can also change to small ubiquitin-like modifier (SUMO)-N and SUMO-C, another
affinity pair. Multiple affinity zipper tests can reduce the conformational bias of the
phosphorylation bought about by the zipper in this design. Using our newly developed
enrichment and analysis techniques, we can investigate all the phosphorylated protein samples
and pool the data together, hoping to reveal specific trends in phosphorylation that will be
beneficial for the design and development of therapeutic drugs that inhibit or reverse the
phosphorylation on Alzheimer tau. [5-6]

186

His6-Fos-GSK

Phosphorylation

Jun-Tev-Tau

Figure 6.3. Scheme of the protein interaction module-assisted phosphorylation. The leucine
zipper Fos and Jun bring the enzyme and substrate together after they are co-produced in cells,
and phosphorylation occurs in vivo.

6.2.3 De novo sequencing of monoclonal antibodies using limited proteolysis
and a middle-down approach
De novo monoclonal antibody sequencing is drawing great interest from many companies
because the obtained sequence can be directly used for recombinant protein production. High
resolution MS instrumentation with top-down strategies make de novo approaches possible.[7]
However, direct, efficient fragmentation of a protein larger than 150 kD is not sufficient enough
to resolve single amino acid residues.[8] The common strategy is to cut the protein then analyze
the parts, and finally map the information together. Papain is frequently used to cut antibodies
because it separates the fragment antigen-binding (Fab) of immunoglobulin G (IgG) from the
fragment crystallizable region (Fc). Another method to cut the antibody is to dissociate the heavy
chains from the light chains by breaking the disulfide bonds linking them.[9] Furthermore, size
exclusion chromatography (SEC) can separate the heavy and light chains.[10] According to Prof.
Gavin Reid’s suggestion, membrane digestion of purified antibody light chain using different
residence times combined with de novo sequencing of the large peptides may provide
comprehensive sequence information. Subsequent mapping and integration of those data should
187

reveal the light chain sequence. Research in this area could begin with commercially available
IgG from Sigma to optimize the reduction (breaking the disulfide bonds between heavy and light
chains) efficiency and separation conditions for SEC in terms of elution buffer, flow rate and the
amount of starting material. Such research will benefit from collaboration with a group equipped
with a fast protein liquid chromatography system to perform the separation. Some preliminary
trials of IgG reduction and separation suggest that the superdex 200 10/300 GL column from GE
Healthcare separates proteins with masses similar to heavy and light chains. Denaturing or
disulfide bond reforming during the separation will influence the separation, so the elution buffer
also must be optimized. Similarly, membrane digestion with MS-analysis requires optimization
with the model IgG protein. Success in de novo antibody sequencing using the limited membrane
digestion approach will greatly increase the impact of protease-modified membranes.

6.2.4 Functional membranes for removal of affinity tags
Recombinant proteins often incorporate affinity tags at a terminus to enhance the protein
solubility in some cases and also facilitate the detection and purification of the protein.[11]
Invented by Roche Corporation, the polyhistidine-tag (His-tag) is the most common tag for
purifying a target protein from a whole cell lysate.[12] After the protein purification, the His-tag
is often removed to avoid its potential influence in down streaming analysis or protein usage. In
many cases, the His-tag is attached to a protein via a cleavable linkage of artificial sequence or
protein, and removed by tobacco etch virus (TEV), thrombin or SUMO proteases. Specifically,
the Tev protease recognizes a seven amino acid sequence (Glu-Asn-Leu-Tyr-Phe-Gln-Gly) and
cleaves between Gln-Gly; the thrombin recognizes the sequence (Leu-Val-Pro-Arg-Gly-Ser) and
cleaves between Arg and Gly; and the SUMO protease directly recognizes the tertiary structure
of SUMO protein and removes the SUMO. Removal of these linkers also removes the His188

tag.[13] Our group has extensive experience in protein purification using Ni-nitrilotriacetic acid
(NTA)-containing membranes,[14] as well as immobilization of proteases for efficient
digestion.[15] Thus, we should be able to immobilize the TEV, Thrombin or SUMO proteases
onto a membrane for tag removal, and flow of the cleaved mixture through a second membrane
can capture the cleaved tag to yield purified protein. Although the traditional protocol is widely
used in many labs (Scheme 6.1), the high enzyme density in a membrane may increases the
digestion efficiency and decreases the time, and the immobilization could stabilize the protease
and allow its reuse.

189

Tag-Removal Digestion
Traditional Workflow
Ni-NTA His6-Linker-Protein

NM

Imidazole
Elution
Imidazole
Removal
Solution
Cleavage
Tag and
Protease
Recapture

Membrane Workflow
Size exclusion Membrane (SM)
Ni-NTA Membrane (NM)
Protease Membrane (PM)

Imidazole
Elution
Spin column

His6-Protease

SM
Imidazole
Removal

PM

Membrane
Tag-Removal
NM

Tag
Recapture
Protein

Protein

Scheme 6.1. Comparison of enzymatic tag-removal in traditional and membrane-based work
flows. In the traditional workflow, imidazole is applied to elute the fusion protein from the NiNTA beads, and before the in-solution tag cleavage, the imidazole is removed via spin column to
facilitate the later tag recapture step. The Ni-NTA beads recapture the His6 tag and His6-protease
and leave the pure target protein. In contrast, the proteins are eluted from Ni-NTA membrane,
and size-exclusion membrane is used to capture the imidazole. Then the protein will undergo
membrane digestion to remove the His6-tag, which will be recaptured by another Ni-NTA
membrane. Pure proteins will be in the last flow-through.
190

6.3

Summary of Future Work
The proposed future work has three directions. The first is the use of TiO2-modifed

membranes combined with limited digestion and intact protein analysis for tau phosphorylation
mapping. This should provide both phosphorylation sites and stoichiometry information to
correlate with biological consequences. The second direction is de novo monoclonal antibody
sequencing using membrane digestion combined with MS analysis. If successful this approach
would provide a routine method for de novo antibody sequencing, an extremely attractive but
unrealized area. The last direction is the development of membrane-based methods to remove the
affinity tag from a recombinant protein. Compared to traditional in-solution cleavage, the
membrane tag removal could provide an alternative method with higher efficiency and less
digestion time.

6.4

Acknowledgement
We are grateful to the U.S. National Institutes of Health (CHE-1152762 to M.B.) for

funding this work.

191

REFERENCES

192

6.5

References

1.

Hanger, D. P.; Noble, W., Functional Implications of Glycogen Synthase Kinase-3Mediated Tau Phosphorylation. Int. J. Alzheimers Dis. 2011, 2011, 352805.

2.

Kellie, J. F.; Tran, J. C.; Lee, J. E.; Ahlf, D. R.; Thomas, H. M.; Ntai, I.; Catherman, A.
D.; Durbin, K. R.; Zamdborg, L.; Vellaichamy, A.; Thomas, P. M.; Kelleher, N. L., The
Emerging Process of Top Down Mass Spectrometry for Protein Analysis: Biomarkers,
Protein-Therapeutics, and Achieving High Throughput. Mol. Biosyst. 2010, 6 (9), 15321539.

3.

Siuti, N.; Kelleher, N. L., Decoding Protein Modifications Using Top-Down Mass
Spectrometry. Nat. Methods 2007, 4 (10), 817-821.

4.

Voss, K.; Gamblin, T. C., GSK-3beta Phosphorylation of Functionally Distinct Tau
Isoforms Has Differential, but Mild Effects. Mol. Neurodegener. 2009, 4 (1), 18.

5.

Hong, M.; Chen, D. C. R.; Klein, P. S.; Lee, V. M. Y., Lithium Reduces Tau
Phosphorylation by Inhibition of Glycogen Synthase Kinase-3. J. Biol. Chem. 1997, 272
(40), 25326-25332.

6.

Mazanetz, M. P.; Fischer, P. M., Untangling Tau Hyperphosphorylation in Drug Design
for Neurodegenerative Diseases. Nat. Rev. Drug Discovery 2007, 6 (6), 464-479.

7.

Resemann, A.; Wunderlich, D.; Rothbauer, U.; Warscheid, B.; Leonhardt, H.; Fuchser, J.;
Kuhlmann, K.; Suckau, D., Top-Down De Novo Protein Sequencing of a 13.6 Kda
Camelid Single Heavy Chain Antibody by Matrix-Assisted Laser Desorption IonizationTime-of-Flight/Time-of-Flight Mass Spectrometry. Anal. Chem. 2010, 82 (8), 3283-3292.

8.

Bondarenko, P. V.; Second, T. P.; Zabrouskov, V.; Makarov, A. A.; Zhang, Z., Mass
Measurement and Top-Down HPLC/MS Analysis of Intact Monoclonal Antibodies on a
Hybrid Linear Quadrupole Ion Trap–Orbitrap Mass Spectrometer. J. Am. Soc. Mass
Spectrom. 2009, 20 (8), 1415-1424.

9.

Gagnon, P.; Rodriquez, G.; Zaidi, S., Dissociation and Fractionation of Heavy and Light
Chains from Igg Monoclonal Antibodies. J. Chromatogr. 2011, 1218 (17), 2402-2404.

193

10.

Liu, H.; Gaza-Bulseco, G.; Chumsae, C., Analysis of Reduced Monoclonal Antibodies
Using Size Exclusion Chromatography Coupled with Mass Spectrometry. J. Am. Soc.
Mass Spectrom. 2009, 20 (12), 2258-2264.

11.

Waugh, D. S., An Overview of Enzymatic Reagents for the Removal of Affinity Tags.
Protein Expr. Purif. 2011, 80 (2), 283-293.

12.

Hochuli, E.; Bannwarth, W.; Dobeli, H.; Gentz, R.; Stuber, D., Genetic Approach to
Facilitate Purification of Recombinant Proteins with a Novel Metal Chelate Adsorbent.
Nat. Biotechnol. 1988, 6 (11), 1321-1325.

13.

Arnau, J.; Lauritzen, C.; Petersen, G. E.; Pedersen, J., Current Strategies for the Use of
Affinity Tags and Tag Removal for the Purification of Recombinant Proteins. Protein
Expr. Purif. 2006, 48 (1), 1-13.

14.

Jain, P.; Vyas, M. K.; Geiger, J. H.; Baker, G. L.; Bruening, M. L., Protein Purification
with Polymeric Affinity Membranes Containing Functionalized Poly(Acid) Brushes.
Biomacromolecules 2010, 11 (4), 1019-1026.

15.

Xu, F.; Wang, W. H.; Tan, Y. J.; Bruening, M. L., Facile Trypsin Immobilization in
Polymeric Membranes for Rapid, Efficient Protein Digestion. Anal. Chem. 2010, 82 (24),
10045-10051.

194