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RELATIONSHIP BETWEEN GLYCOLYTIC OSCILLATIONS
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presented by

NICHOLAS L. BERRY

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RELATIONSHIP BETWEEN GLYCOLYTIC OSCILLATIONS AND PORK COLOR
AND WATER-HOLDING CAPACITY

By

Nicholas L. Berry

A THESIS
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

MASTER OF SCIENCE
Department of Animal Science

2003

ABSTRACT

RELATIONSHIP BETWEEN GLYCOLYTIC OSCILLATIONS AND PORK COLOR
AND WATER-HOLDING CAPACITY

By Nicholas L. Berry

The objective of this study was to determine if differences exist in the pattern of
glycolysis in muscle extracts from superior and inferior quality pork loins.
Characteristics of superior (n=6) and inferior (n=6) quality loins, respectively, were as
follows: 45 min pH (6.40 i 0.06 vs 5.92 i 0.10), percent fluid loss alter centrifugation
(10.22 :t: 0.47 vs 20.70 :t: 0.32), percent drip-loss by the suspension method (0.66 :i: 0.08
vs 3.23 :t: 0.45), and day l L“ (51.37 i 0.66 vs 56.68 :i: 0.86). Longissimus muscle
samples were obtained at 20 min postmortem and were immediately frozen in liquid
nitrogen and stored at -80° C until analysis. Samples were homogenized in 90 mM
potassium phosphate, pH 6.5, and 180 mM potassium chloride. A high-speed supernatant
fluid (85,000 x g) was gel-filtered using Sephadex G-25 resin. Gel-filtered muscle
proteins were used to determine in vitro patterns of glycolysis. Aliquots of glycolytic
reaction mixture were removed every 2 min for 46 min and acidified to halt enzyme
activity. Enzymatic assays were used to quantify concentrations of ADP, ATP, and
lactate. An oscillatory pattern of glycolysis was observed using extracts from both
superior and inferior quality pork. No differences in the average ATPzADP ratio or the
overall mean concentrations of lactate or adenine nucleotides were observed in reactions
using extracts from superior and inferior quality samples (P > 0.05). Thus, sarcoplasmic
protein extracts do not appear to produce distinct patterns of glycolysis that are associated
with differences in pork quality. However, this system will permit identification of

specific metabolites that cause differences in the pattern of postmortem glycolysis.

ACKNOWLEDGEMENTS

I would like to begin my acknowledgements by thanking my friend and major
professor, Dr. Matt Doumit. I have truly enjoyed the Opporttmity to further my education
under your guidance at Michigan State. Thank you very much for being patient and for
giving me the chance to prove myself as a student, a researcher, and a person.

Thank you Dr. Al Booren, Dr. Gretchen Hill, and Dr. Gale Strasburg for serving
on my committee. I greatly appreciate your guidance and support throughout my stay at
MSU. I have enjoyed getting to know each of you, not only as researchers, but more
importantly as friends.

Thank you Chuck Allison and Emily Helman for not only your time and hard
work, but also your fi'iendship. To Chuck, I am greatly appreciative of your efforts to
improve my skills as a researcher. You have been a great role model in the lab, as well as
an excellent friend to me throughout my time in East Lansing. To Emily, I am greatly
indebted to you for your time and patience while developing my skills in the lab. You
have been an excellent teacher and friend throughout my stay at MSU.

I would also like to extend my thanks to the undergraduates that have worked in
the Doumit lab throughout the completion of my project. I have enjoyed your friendship,
and am greatly appreciative of the time and effort you have put forth.

And last, but certainly not least, thanks to all of my friends and family. To my
parents, thank you for encouraging me to always pursue my dreams and for instilling in
me the value of hard work and persistence. To Kohl Schrader, thank you for putting up
with me alter long hours in the lab, and for always putting a smile on my face. I’m

greatly appreciative of all your support.

iii

TABLE OF CONTENTS

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

LIST OF TABLES v
LIST OF FIGURES vi
LIST OF ABBREVIATIONS vii
INTRODUCTION 1
LITERATURE REVIEW 3
A. Pork Quality 3

B. Genetic Basis for PSE Pork 6

C. Biochemical and Physical Aspects of PSE Pork 7

D. Glycogen Storage in Skeletal Muscle 8

E. Anaerobic Metabolism of Glucose 9

F. Postmortem ATP-utilization 10

G. Regulation of Glycolysis 11

H. Glycolytic Oscillations 13

I. Regulators of Glycolysis 15
CHAPTER 1 16
A. Abstract 16

B. Introduction 17

C. Materials and Methods 19

1. Animal and Meat Quality Data Collection 19

2. Preparation of Muscle Protein Extracts 20

3. Polyacrylamide Gel Electrophoresis (PAGE) of Gel-filtered Protein ,,,,,,,,,,,,, 22

4. Myofibril Preparation 22

5. Glycolytic Reaction Conditions 23

6. ATP Assay 24

7. ADP Assay 25

8. Lactate Assay 26

D. Results and Discussion 28

E. Implications 35
APPENDICES 36
LITERATURE CITED 51

 

iv

LIST OF TABLES
Table 1 Means and standard deviations for carcass quality data 36

Table 2 Overall mean concentrations of adenine nucleotides and lactate, and calculated
ATPzADP ratio 36

LIST OF FIGURES

 

 

 

 

 

 

 

Figure 1 Gel-Filtration Protein Elution 37
Figure 2 Polyacrylamide Gel Electrophoresis (PAGE) of Muscle Protein ,,,,,,,,,,,,,,,,,,,, 3 8
Figure 3.1 Adenine nucleotide (ADP and ATP) and lactate concentration of a superior
quality Duroc pig 39
Figure 3.2 Adenine nucleotide (ADP and ATP) and lactate concentration of a superior
quality Pietrain pig 40
Figure 3.3 Adenine nucleotide (ADP and ATP) and lactate concentration of an inferior
quality Duroc pig 41
Figure 3.4 Adenine nucleotide (ADP and ATP) and lactate concentration of an inferior
quality Pietrain pig 42
Figure 3.5 Adenine nucleotide (ADP and ATP) and lactate concentration of a superior
quality Pietrain pig 43
Figure 3.6 Adenine nucleotide (ADP and ATP) and lactate concentration of an inferior
quality Duroc pig 44
Figure 4 Superior vs Inferior Quality - lactate accumulation firm 0 to 46min ,,,,,,,,,,,,,, 4 5

Figure 5 In vitro pH decline of superior and inferior quality samples during reactions
from 0 to 46 min 46

Figure 6 Adenine nucleotide (ADP and ATP) and lactate concentration of rat hind limb
muscle 47

Figure 7.1 Lactate accumulation of a supeior quality Duroc pig with the addition of three
differing concentrations of myofibrils from O to 46 min 48

Figure 7.2 Lactate accumulation of an inferior quality Duroc pig with the addition of
three differing concentrations of myofibrils from 0 to 46 min 49

Figure 8 Lactate concentration of pig muscle extract exposed to differing
concentrations of citrate from 0 to 46 min 50

vi

LIST OF ABBREVIATIONS
ADP - Adenosine Diphosphate
AMP - Adenosine Monophosphate
APS - Ammonium Persulphate
ATP - Adenosine Triphosphate
CWHC - Centrifugal Water Holding Capacity
DFD - Dark, Firm and Dry
DRIPl - Suspension Drip Measured from 24 to 48 hours postmortem
EDTA - Ethylenediaminetetraacetic Acid
G-6-P - GIucose—6-Phosphate
GTP - Guanosine Triphosphate
HCl - Hydrochloric Acid
HK - Hexokinase
KOH - Potassium Hydroxide
L1 - Day 1 L“
LM - Longissirnus Muscle
MSU - Michigan State University
NAD - Nicotinamide Adenine Dinucleotide
NADH - Nicotinamide Adenine Dinucleotide, reduced form
NPPC - National Pork Producer’s Council
Pi - Inorganic Phosphate
PFK - Phosphofi'uctokinase

PK - Pyruvate Kinase

vii

PMSF - Phenylmethylsulfonylfluoride

PSE - Pale, Soft and Exudative

PSS - Porcine Stress Syndrome

RFN - Red, F inn and Non-exudative

RN' - Rendement Napole

RYRI — Skeletal Muscle Ryanodine Receptor

SDS - Sodium Dodecyl Sulfate

SDS-PAGE - Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis
SR - Sarcoplasmic Reticulum

TEMED - N,N,N’,N’- Tetramethylenediamine

WHC - Water-holding Capacity

viii

INTRODUCTION

Today’s U.S. pork industry is experiencing phenomenal growth as it continues to
meet consumer demand. Pork is the most widely eaten meat worldwide. Consumption of
pork in the United States has proven to be a valuable protein source as Americans
consume 66 lbs/person/year on a carcass disappearance basis (USDA, 2002 Agricultural
Statistics). Pork production in the United States is a vital part of the economy. Nearly 19
billion pounds of pork, with a retail value of $38 billion, was processed from about 97
million hogs in 2001. The US. pork industry is responsible for over $72 billion in total
domestic economic activity. In addition, the pork industry adds over $27 billion of value
to basic production inputs such as corn and soybeans (NPPC, 2003).

Currently, the United States is one of the world’s leading pork-producing
countries and is second to only Canada as the largest exporter in the world. Pork
production in the US. accounts for about 10% of the world’s supply. Additionally, there
are approximately 85,760 pork operations today, as compared to nearly three million in
the 19508 (NPPC, 2003). The aforementioned statistics mirror the changes seen within
the industry, and illustrate the emphasis placed on production and efficiency.
Subsequently, the emphasis placed on lean, fast growing pigs has compromised pork
quality. Incidence of pale, soft, and exudative (PSE) pork has risen from 10% to 15.5%
between 1996 and 2003 (Cannon et al., 1996; Stetzer and McKeith, 2003). Additionally,
inferior quality problems cost the industry an estimated $90 million annually (Stetzer and
McKeith, 2003).

Developing a thorough understanding of the biochemical processes that are

integral in explaining variation observed in ultimate pork quality is of utmost concern.

The objective of this study is to determine if differences exist in the pattern of glycolysis
in muscle extracts from superior and inferior quality pork loins. Collectively, these data
will provide a better understanding of glycolytic behavior during the postmortem time
period. Moreover, this system will allow for identification of specific regulators that
cause difl‘erences in the pattern of postmortem glycolysis. This knowledge could lead to
manipulation of live pigs or carcasses by altering events that influence postmortem

energy utilization.

LITERATURE REVIEW
Pork Q_u_ah'g

Red, firm and non-exudative (RFN) pork is considered to be the ideal pork
product desired by consumers based on both color and textural attributes. The National
Pork Producers Council (NPPC) Pork Quality Solutions Team established pork quality
targets in order to define the ideal quality benchmark (NPPC, 1998). The acceptable
subjective color score ranges fiom 3 to 5 on a 6 point scale (1 = pale pinkish gray to
white and 6 = dark purplish red). Acceptable lightness (L‘) is measured objectively
using a Minolta colorimeter and ranges from 37 to 49. Ultimate pH should be between
5.6 and 5.9. Additionally, ideal pork should have less than 2.5% drip loss when
measured at 24 hours postmortem (NPPC, 1998). RFN pork exhibits the aforementioned
attributes desired by the quality benchmarking system.

A limited pH decline with the onset of rigor mortis at a relatively high pH (2 6.0)
will produce meat exhibiting a dark red color, a firm texture and a dry appearance. Dark,
firm and dry (DFD) pork is often associated with long-term stress (i.e. prolonged injury,
illness, or extended hauling) that results in depletion of muscle glycogen prior to harvest.
Subsequently, less lactate and IT ions accumulate, and the ultimate pH is higher (2 6.0).
Because of this higher pH, proteins in DFD meat have a relatively high net negative
protein charge and undergo minimal protein denaturation (Briskey et al., 1966). These
factors result in relatively high water-holding capacity (WHC) due to greater repulsion
between myofilaments and greater protein/water interaction compared to RFN or PSE
pork. However, the higher ultimate pH of DFD pork makes this product more susceptible

to spoilage by bacteria and subsequently shortens shelf life. In addition, many consumers

discriminate against DFD pork due to its unacceptable dark purple color (Kauifinan et al.,
1 992).

Pale, soft and exudative pork is characterized by its pale color and inferior WHC.
The product is very watery in appearance and flaccid in texture. As selection for leaner,
heavier weight pigs with maximized muscularity has occurred, so has the increased
occurrence of PSE pork (Lonergan et al., 2001). A survey consisting of fourteen packing
plants revealed that approximately 16% of pork produced exhibited PSE characteristics
(Kauffrnan et al., 1992). The Pork Quality Chain Audit (Cannon et al., 1996) and the
National Pork Producers Council (NPPC 1991), also found that greater than 10% of all
pork carcasses generated in the US. contained PSE meat. A loss of thirty-five cents per
carcass is attributable to WHC problems associated with PSE (Cannon et al., 1996).
Thus, PSE costs the pork industry millions of dollars every year (Stetzer and McKeith,
2003).

PSE pork characteristics are hard to induce in pigs that are less susceptible to
stress suggesting that there is a complex physiological and genetic difference among
genetic lines that causes some pigs to have a greater predisposition to develop an
abnormal pH decline (Gerrard, 1997). Muscles considered to be PSE generally have
significantly lower pH values and higher muscle temperatures at forty minutes
postmortem (Briskey et al., 1959). Approximately 20% of the sarcoplasmic and
myofibrillar proteins are denatured as a result of rapid pH decline accompanied by high
carcass temperatures (Honikel and Kim, 1986). Increased lightness has been associated
with reduced sarcoplasmic protein solubility, and to a lesser extent myofibrillar protein

solubility (Joo etal., 1999). Phosphorylase, creatine kinase, triose phosphate isomerase,

and myokinase are sarcoplasmic proteins shown to precipitate onto myofibrils in PSE
pork (Joo et al., 1999). Reduced WHC has been attributed to denaturation of
sarcoplasmic (Joo et al., 1999; Wilson and van Laack, 1999) and myofibrillar proteins
(Offer, 1991; Warner et al., 1997). Breaks in the cell membrane through which fluid can
quickly exude from the cell are also a characteristic of PSE muscle (Honikel and Kim,
1986).

The most undesirable aspect of PSE pork is the reduced WHC. The inability of
meat to bind water greatly reduces the opporttmities the packer has to enhance the
product and has a detrimental effect on consumer acceptability. For example, during
storage PSE hams lose three-fold more moisture than normal hams and seven-fold more
moisture than DFD hams (Kauffrnan et al., 1978). Moisture losses during chilling, as
well as during curing and smoking, are higher in PSE hams than in normal or DFD hams.-
Harn and shoulder cuts that are severely PSE exhibit 33% and 11% lower WHC,
respectively, when compared with normal barns and shoulders (Cannon et al., 1995).

Kauffman .et al., (1999) subjected eight DFD pork loins and eight PSE pork loins
to a series of objective and subjective measures to demonstrate meat quality. When
compared to PSE loins, DF D loins averaged 1.5 units higher ultimate pH, 4.7% less drip
loss and 13.6% more bound water. Additionally, DFD loins were darker, firmer and
more tender. Not all of the previously mentioned attributes allow for optimum quality of
pork. DF D meat is a prime target for bacterial problems as many of the species that grow
on meat survive within a pH range of 5.0 to 8.0, with optimum growth occurring around

pH 7.0 (Cannon et al., 1995). As an industry, our responsibility is to maximize the

quality of the final meat product purchased by the consumer. This goal can be achieved
with a further understanding of biochemical events that occur within the muscle tissue.
Genetic Basis for PSE Pork

Porcine stress syndrome (PSS), also referred to as malignant hyperthermia, is a
genetic abnormality characterized by the inability of the pig to adapt to stressors. F ujii et
a1. (1991) demonstrated that PSS was associated with a genetic mutation (HAL-1843) in
the muscle ryanodine receptor (RYRl; calcium-release channel), which was associated
with abnormal leakage of calcium from the channel. Rapid ATP utilization and
glycolysis, such as that induced by excessive calcium release through defective RYRl,
are often associated with development of PSE pork (Honikel et al., 1986). Stimulation of
muscle contraction, sarcoplasmic reticulum (SR) calcium ATPase (calcium pump)
activity, and glycolysis by sarcoplasmic calcium results in heat production and muscle
pH decline. Consequently, the combination of low pH and high temperature early
postmortem results in a greater occurrence of protein denaturation in pigs with this
condition as compared to stress-resistant genotypes (Briskey et al., 1966; Louis et al.,
1993)

Based on the findings of F ujii et a1. (1991), a DNA-based test was established that
could distinguish between homozygous carrier, heterozygous carrier, and homozygous
normal pigs with respect to the HAL-1843 mutation. This genetic test largely replaced
halothane gas testing of swine, and efforts were made by the National Pork Board to
eliminate the HAL-1843 mutation from commercial populations. Nonetheless, Murray
and Johnson (1998) reported that of a population of 1006 pigs in two packing plants, 90%

of the PSE condition was caused by factors other than the HAL-1843 gene.

The rendement napole (RN-) gene was recently identified as a defect in the AMP-
activated protein kinase (Milan et al., 2000). The dominant RN- allele results in higher
than normal muscle glycogen stores and an extended postmortem pH decline that leads to
pork with a lower than normal ultimate meat pH, higher reflectance (lighter meat color),
reduced WHC, and dramatically reduced carcass yield (LeRoy et al., 2000). Monin and
Sellier (1985) referred to this condition as the Hampshire effect, due to its prevalence in
the Hampshire breed. Other factors, such as enzyme stability at lower pH, must play a
vital role in the continuation of glycolytic metabolism in these animals.

Biochemical and Physical Aspects of PSE Pork

Several factors influence the WHC of pork, including myofibrillar lattice spacing,
cytoskeletal links, membrane permeability, and the size of fluid channels in the
extracellular space (Purslow et al., 2001; Honikel, 2002). Development of PSE pork
characteristics can result from a rapid antemortem and/or postmortem pH decline, or an
extended postmortem pH decline. The rapid drop in pH in PSE muscles within the first
hour after slaughter is influenced by a high ATP turnover. The increased turnover in
ATP is attributable to the release of Ca2+ ions that induce muscle contraction, and
subsequently ATP utilization. Glycogenolysis is also stimulated by conversion of
phosphorylase b into phosphorylase a via phosphoylase kinase, which is activated by
Ca2+ ions (Honikel, et al., 1986). Anaerobic glycolysis results in production of ATP,
lactate and hydrogen ions. Hydrogen ion accumulation lowers the muscle pH. Relatively
low pH combined with high muscle temperature during the early postmortem period,
causes denaturation and reduced solubility of sarcoplasmic proteins (Sayre and Briskey,

1963; Scopes, 1964; 100 et al., 1999) and myosin (Offer, 1991; Warner et al., 1997).

Pale color and reduced WHC associated with PSE pork have been primarily
attributed to denaturation of sarcoplasmic and myofibrillar proteins, respectively (Joo et
al., 1999). However, Wilson and van Laack (1999) demonstrated that when myofibrils
from either PSE or normal pork were combined with sarcoplasmic extract from PSE
meat, the water-holding capacity of the myofibrils was lower than when combined with
extract from normal pork. These authors concluded that sarcoplasmic proteins also
influence WHC, through as yet undefined mechanisms. One possibility is that denatured
sarcoplasmic proteins adsorb onto the surface of myofibrils, thereby shielding the
charged groups available for fluid binding (Bendall and Wismer-Pedersen, 1962; Boles et
al., 1992).

Glycogen Storage in Skeletal Muscle

Glycogen is the storage form of glucose within skeletal muscle. Glycogen is
made up of oil-4 and (ll-6 linkages of glucose molecules. The synthesis of glycogen
involves a protein primer, glycogenin, which acts as a hexosyltransferase (Alonso et al.,
1995). Glycogen synthesis begins as the amino acid tyrosine-194 on the glycogenin
residue and becomes autocatalytically glucosylated by the addition of UDP-glucose into
the system (Lomako et al., 1993). On average, seven other glucose residues are added to
tyrosine-194 in the same manner. These eight glucose residues become the primer for the
synthesis of proglycogen. Proglycogen is a stable intermediate for both the synthesis and
breakdown of glycogen (Alonso et al., 1995). This stable intermediate is formed by the
addition of proglycogen synthase and branching enzyme to the fully glycosylated
glycogenin molecule. Proglycogen is then converted to macroglycogen, the storage form

of glucose, by using macroglycogen synthase and branching enzyme. This reaction is

reversible, allowing for the breakdown of macroglycogen into proglycogen by use of
phosphorylase and debranching enzyme (Lomako et al., 1993). In terms of metabolic
activity, the larger macroglycogen (107 D) is inactive while proglycogen (400,000 D) is
an active intermediate (Alonso et al., 1995). In order for glycogen to be used in
metabolism, macroglycogen must first be converted into proglycogen.

The implication of this glycogen storage mechanism is that glycogen storage may
slow down the rate of glycolysis by storing more glucose as the inactive macroglycogen
as opposed to the metabolically active proglycogen. In a study done by Huang and others
(1999), it was observed that when rats were exposed to minimal levels of stress,
proglycogen was exclusively depleted and was also the only molecule that underwent
synthesis and degradation when the rate of glycogen turnover was low. As the stress
levels increased, macroglycogen was recruited in glycogen turnover. This study also
observed that injecting high levels (greater than 10 mU/min") of the hormone insulin
promoted the storage of glucose in the form of macroglycogen as opposed to
proglycogen.

Anaerobic Metabolism of Glucose

Glycolysis is an important pathway for providing energy hour the stepwise
degradation of glucose. It is primarily an anaerobic form of metabolism and its use in
living animals is to produce short-term energy when oxygen is limiting to the organism.
Glycolysis consists of two phases (Garrett and Grisham, 1999). The first phase is a series
of five reactions in which glucose is broken down to two molecules of glyceraldehyde-3-
phosphate. This phase consumes 2 molecules of ATP, or 1 molecule of ATP if glycogen

is utilized as substrate. The second phase converts the two molecules of glyceraldehyde-

3-phosphate into two molecules of pyruvate. The second phase results in a net gain of
four ATP, thus glycolysis results in a net gain of two or three ATP, depending on the
source of glucose.

The conversion of muscle to meat begins with exsanguination. Biochemical
reactions that occur following exsanguination are largely responsible for the ultimate
quality of meat. Loss of blood ceases the exchange of oxygen between the circulatory
system and the muscle tissue. As oxygen is depleted, a shift from aerobic metabolism to
anaerobic metabolism occurs to meet the demands for postmortem ATP utilization.
Anaerobic metabolism results in a less efficient production of ATP and accumulation of
lactate and hydrogen ions, the end products of anaerobic glycolysis (Goll et al., 1984). In
animal tissues that are experiencing anaerobic conditions, the two molecules of pyruvate
that are produced by glycolysis are reduced to lactate by lactate dehydrogenase.
Accumulation of hydrogen ions results in the reduction in pH of pre-rigor muscle as
muscle is converted to meat. The rate of pH decline is related to the temperature of the
muscle. A higher muscle temperature will result in a faster rate of pH decline (Sayre and
Briskey, 1963 ), whereas a lower muscle temperature will result in a slower rate of pH
decline. Once the ATP is completely consumed, and no more is being produced,
permanent crossbridges are formed between myosin and actin to create actomyosin. This
condition is termed rigor mortis.

Postmortem ATP-utilizgtion

The major sites of ATP utilization are the myofibrillar (myosin) ATPase, which is

activated during muscle contraction, and the calcium ATPase pump that functions to

resequester calcium into the sarcoplasmic reticulum (Bechtel and Best, 1985). Creatine

IO

phosphate is readily available for regeneration of ATP via a reaction catalyzed by the
enzyme, creatine phosphokinase. Generation of ATP also occurs by a reaction catalyzed
by myokinase, which converts two molecules of ADP to ATP and AMP (Bechtel and
Best, 1985). These reactions may have a sparing effect on the rate of postmortem
glycolysis. However, both myokinase and creatine phosphokinase are susceptible to
denaturation as acidic postmortem conditions develop (Joo et al., 1999). Thus, the rate of
glycolysis increases in response to increased ATP utilization.

mum of Glycolysis

In living skeletal muscle, energy utilization and energy production are highly
coordinated events. In fact, Conley et al. (1997) suggested that elevated sarcoplasmic
calcium activates muscle contraction, glycogenolysis and glycolysis in parallel, since
glycolytic rate is dependent on muscle stimulation frequency and independent of ADP,
AMP and P, concentrations. Once again, this highlights the importance of calcium
regulation on muscle metabolism. Control over glycolytic flux, or the flow of
intermediates through glycolysis, may also be controlled by covalent modification
(enzyme phosphorylation and dephosphorylation), substrate control, and allosteric control
mediated by changes in metabolite and co-factor concentrations (reviewed by Connett
and Sahlin, 1996). The regulation of glycolytic flux is complex.

Phosphofructokinase (PF K) is generally regarded as the primary regulatory
enzyme of glycolysis. It was on this premise that Sayre et al. (1963) investigated PFK
activity in porcine muscle extracts. These authors determined that PFK and
phosphorylase activities were not associated with the rate of pH decline in longissimus

muscle of Hampshire, Poland China and Chester White pigs. Surprisingly, Allison et al.

11

(2003) found that maximal in vitro PFK activity extracted from longissimus samples
fiozen at 20 minutes postmortem was inversely correlated with loin chop fluid loss. This
likely results from early postmortem inactivation of the acid labile PFK enzyme in
muscle undergoing rapid glycolysis.

Although glycolytic enzyme capacity does not appear to explain a large
proportion of the variation in pork color and WHC, other levels of glycolytic regulation
may contribute to the rapid pH decline often associated with PSE pork. A detailed
understanding of factors that regulate postmortem glycolysis in porcine skeletal muscle
will improve efforts to control the rate of muscle pH decline.

The reversible binding of enzyme-enzyme and enzyme-contractile protein
interactions provide additional possibilities for the regulation of glycolytic flux in skeletal
muscle. The proportions of several glycolytic enzymes bound to contractile proteins
increase with increased rates of glycolysis, and this may provide a mechanism for
enhancing metabolite transfer rates (Parkhouse, 1992). Lee et al. (1989) summarized
several studies demonstrating that PFK is phosphorylated in contracting muscle when the
need for energy is high. Modification of PFK by phosphorylation favors enzyme binding
to actin by increasing its apparent affinity for F-actin. It is unclear if postmortem
conditions induce enzyme modifications that are similar to those that occur under normal
physiological conditions.

How (or it) the functional coupling of glycolytic enzymes to the major sites of
energy utilization (myosin ATPase and calcium ATPase) affect pork quality is currently

unknown. However, the effects of this coupling may influence glycolytic rate, ultimate

12

enzyme location and the degree of denaturation of sarcoplasmic proteins, which in turn,
may influence color (Joo et al., 1999) and WHC of pork (Wilson and van Laack, 1999).

Kastenschmidt et al. (1968) quantified levels of glycolytic intermediates and co-
factors in longissimus muscles that exhibited fast and slow rates of postmortem
glycolysis. Muscles were classified as either “fast-glycolyzing” or “slow-glycolyzing”
based on the rate of their post-mortem decline in pH. If muscle pH declined to 5.5 or
below at 30 minutes post-mortem, the muscle was termed “fast-glycolyzing.” Contrary
to this, if muscle pH was 6.0 or higher at 60 minutes post-mortem, the muscle was termed
“slow-glycolyzing.” “Fast-glycolyzing” muscles are in a highly anaerobic state prior to
or simultaneous with death.

Results of this thorough investigation led the authors to conclude that accelerated
glycolytic rates appear to result fi'om coordinated stimulation of glycogen phosphorylase,
PFK and pyruvate kinase (PK) enzymes. These enzymes have traditionally been
considered to catalyze rateodetermining steps of glycogenolysis and glycolysis in skeletal
muscle. These reactions are far from equilibrium and are catalyzed by PFK and PK and
proceed with a large decrease in free energy. However, the precise regulation of these
enzymes in postmortem muscle is unclear.

Glycolytic Oscillations

 

For decades, it has been known that the rate of postmortem glycolysis is variable
in the pig muscle (Briskey et al., 1966). Postmortem muscle energy utilization is
monitored by measuring pH at specified times. Continuous pH measurement in muscle is
difficult to achieve with current instrumentation, yet pH decline data are typically drawn

as curvilinear. This measurement undoubtedly reflects the pH of tissue, rather than

13

individual muscle fibers, and may not accurately reflect the nature of postmortem
glycolysis in individual muscle fibers. Additionally, the measurement of early
postmortem pH ofien produces variable results that are not highly correlated with pork
quality traits. Oscillatory behavior of the glycolytic pathway has been well documented
(reviewed by Smolen, 1995; Tomheim, 1979). In cell-flee extracts of skeletal muscle,
glycolytic oscillations are generated by repeated bursts of PF K activity. When the
[ATP]/ [ADP] ratio decreases to a trigger level, this initiates a sudden increase, or burst,
in glycolytic flux that restores a high [ATP]/[ADP] ratio. In postmortem tissue,
glycolytic bursts may also result in rapid and localized acidification, which could
exacerbate protein denaturation.

Under conditions encountered in skeletal muscle, oscillatory behavior of
glycolysis has advantages over steady state behavior for regulation of carbohydrate
utilization and maintenance of a high [ATP]/[ADP] ratio. It also has greater
thermodynamic efficiency (Tomheirn et al., 1979, 1991; Termonia et al., 1981). It is not
known if postmortem skeletal muscle of the pig exhibits oscillatory patterns of glycolysis
and associated bursts in acidification. However, postmortem muscle pH decline may be
sporadic in some pigs (R.C. Johnson and CF. Allison; unpublished data). In whole ,
muscle, glycolytic oscillations may frequently be obscured by cell-to—cell asynchrony
(Tomheirn et al., 1979). Bursts of acidification resulting from early postmortem
glycolytic oscillations would likely cause more extensive protein denaturation than a
gradual pH decline. Thus, glycolytic oscillations would be important determinants of
pork color and WHC. The potential contribution of these oscillations to the development

of PSE pork warrants firrther investigation.

14

Regglators of Glycolysis
Oscillatory behavior of glycolysis involves bursts of PFK activity due to

autocatalytic activation by fructose 1,6-bisphosphate. Fructose 2,6-bisphosphate is an
even more potent activator of PFK and is competitive with fructose 1,6-bisphosphate for
the same binding site on PFK. Tomheim et al., (1988) have shown that addition of
fructose 2,6-bisphosphate blocked glycolytic oscillations, but did not affect lactate
accumulation. Additionally, fructose 2,6-bisphosphate decreased the [ATP]/ [ADP] ratio,
demonstrating the advantage of the oscillatory behavior in maintaining a high energy
state.

Citrate has also been shown to modulate glycolytic oscillations by decreasing
their frequency and delaying their on-set (Tomheim et al., 1991). These effects are due
to inhibition of PFK, such that a lower trigger value of the [ATP]/[ADP] ratio must be
reached before a burst of PFK can be initiated. However, citrate inhibition has little
effect on the amplitude of oscillations or the peak [ATP]/ [ADP] ratio (Tomheim et al.,
1991). In glycolytic oscillations with and without citrate, 50 — 100% more AMP was
required to initiate the burst of PFK activity in the presence of citrate. In other words, the
[ATP]/ [ADP] ratio had to fall to a greater extent in the presence of citrate before PFK

was activated (T omheirn et al., 1991).

15

CHAPTER]

RELATIONSHIP BETWEEN GLYCOLYTIC OSCILLATIONS AND PORK COLOR
AND WATER-HOLDING CAPACITY

ABSTRACT

The objective of this study was to determine if differences exist in the pattern of
glycolysis in muscle extracts fiom superior and inferior quality pork loins.
Characteristics of superior (n=6) and inferior (n=6) quality loins, respectively, were as
follows: 45 min pH (6.40 :l: 0.06 vs 5.92 i 0.10), percent fluid loss after centrifugation
(10.22 i: 0.47 vs 20.70 i 0.32), percent drip—loss by the suspension method (0.66 :l: 0.08
vs 3.23 :l: 0.45), and day 1 L“ (51.37 :h 0.66 vs 56.68 :l: 0.86). Longissirnus muscle
samples were obtained at 20 min postmortem and were immediately frozen in liquid
nitrogen and stored at -80° C until analysis. Samples were homogenized in 90 mM
potassium phosphate, pH 6.5, and 180 mM potassium chloride. A high-speed supernatant
fluid (85,000 x g) was gel-filtered using Sephadex G-25 resin. Gel-filtered muscle
proteins were used to determine in vitro patterns of glycolysis. Aliquots of glycolytic
reaction mixture were removed every 2 min for 46 min and acidified to halt enzyme
activity. Enzymatic assays were used to quantify concentrations of ADP, ATP, and
lactate. An oscillatory pattern of glycolysis was observed using extracts fi'om both
superior and inferior quality pork. No differences in the average ATPzADP ratio or the
overall mean concentrations of lactate or adenine nucleotides were observed in reactions
using extracts from superior and inferior quality samples (P > 0.05). Thus, sarcoplasmic
protein extracts do not appear to produce distinct patterns of glycolysis that are associated
with differences in pork quality. However, this system will permit identification of

specific metabolites that cause differences in the pattern of postmortem glycolysis.

16

INTRODUCTION

The National Pork Benchmarking Audit found that live weight and carcass
muscle percentage have increased, and backfat thickness has decreased over the last ten
years (Stetzer and McKeith, 2003). Furthermore, as selection for leaner, heavier weight,
and more muscular pigs has occurred, so has the increased occurrence of PSE pork
(Lonergan et al., 2001). Despite an abundance of research describing PSE pork
characteristics, and a reduction in the frequency of major genes with known deleterious
effects on pork quality, Cassens (2000) concluded that little progress has been made in
reducing the incidence of PSE pork

Understanding events that dictate superior and inferior quality is essential for
development of new strategies to improve the quality and consistency of pork. Briskey et
al. (1966) have shown that the rate of postmortem glycolysis is variable in pig muscle.
Pale, soft and exudative pork is caused by the denaturation of muscle proteins that result
when carcass muscles experience a low pH and high temperature (Briskey and Wismer-
Pedersen, 1961). However, low early postmortem pH does not explain all PSE seen in
pig muscle. Pork with an apparently normal pH decline may exhibit pale and/or sofi and
exudative characteristics. Observed inconsistencies in pH decline within a muscle may
suggest differences in the pattern of pH decline among muscles or carcasses. In support
of this, a recent report suggests that postmortem muscle pH decline may be sporadic in
some pigs (Johnson, RC, 2001), suggesting that differences may exist in the patterns of
energy metabolism during the postmortem period. Oscillatory behavior of the glycolytic
pathway in vitro has been well documented (Tomheim et a1. 1971, 1973, 1974, 1991). In

whole muscle, glycolytic oscillations may frequently be obscured by cell-to-cell

17

asynchrony (T omheim et al., 1979). We hypothesized that inferior pork water-holding
capacity and pork color are associated with more pronounced glycolytic oscillations,

which exacerbate protein denaturation.

18

MATERIALS AND METHODS
Animal and Meat Min Data Collection

Sires from Duroc and HAL-1843-free Pietrain lines were mated to Yorkshire and
F 1 Yorkshire-Landau gilts. Crossbred progeny were raised in uniform conditions at the
Michigan State University (MSU) Swine Teaching and Research Farm. Four gilts from
each sire group were harvested on each of four days within a 2-week period at the MSU
Meat Laboratory. Since this project required no experimental manipulation of live
animals it was exempt from filing an animal use form based on AUCAUC Policy #9,
Policy on Submission of an Animal Use Form, projects involving research on tissues or
fluids collected from a food animal killed in a slaughterhouse. Pigs were harvested using
standard industry procedures following the Code of Federal Regulations for Humane
Slaughter of Livestock.

Muscle tissue and pork quality data were collected as previously described by
Allison et al. (2003). Briefly, a longissimus muscle (LM) sample was taken midway
between the last rib and the cranial edge of the iliurn on the left side of the carcass 20 min
postmortem. Each sample (~2.5 cm thick) was cut into 0.5 cm3 pieces, frozen in liquid
nitrogen, and stored at -80° C for later use. Longissirnus muscle pH was measured at 20,
45, 180 min, and 22 h postmortem with a portable pH meter (Model 1140, Mettler-
Toledo, Woburn, MA) equipped with a puncture-type combination pH electrode (Lot
406-M6-DXK-57/25, Mettler-Toledo, Woburn, MA). At 24 h postmortem, duplicate 10
g samples were obtained from the cranial edge of the loin section to determine WHC by
high-speed centrifugation (CWHC) at 40,000 x g for 30 min (Honikel and Hamm, 1994).

Two 2.5 cm thick loin chops were removed between the 12m and last rib of the section.

19

These chops were used to determine drip loss (DRIPl) by the suspension method from 24
to 48 h postmortem (Honikel and Hamm, 1994) and color (15 min bloom time; CIE L‘,
D65, 2 degree standard observer and 50 mm orifice) at 24 h postmortem with a Minolta
chromameter (CR-3 10; Ramsey, NJ).

Based on difl‘erences in 45 min pH, CWHC, DRIPl, and L‘, samples from three
Duroc- and three Pietrain-sired pigs with superior meat quality traits, and from three
Duroc- and three Pietrain-sired pigs with inferior meat quality traits were selected. Pigs
that were most extreme in their quality differences were selected for the current
experiment.
man of muscle motein extracts

Chemicals used in the muscle extraction and assays were purchased from Sigma
Chemical Company (St. Louis, MO). Methods used were adapted from those described
for rat muscle by Tomheim et al. (1973, 1974, 1979, 1988, and 1991). Frozen LM
samples (~7 g) were homogenized in 3 volumes of cold homogenization buffer (90 mM
potassium phosphate, pH 6.5, and 180 mM potassium chloride). Homogenization was
performed using a Polytron (Brinkrnan, Westbury, NY) for 3 x 30—sec bursts (setting 4),
with 15 sec between bursts. Samples were continuously cooled with ice during
homogenization. The homogenate was stirred slowly at 4° C for 1 hour, and then
centrifuged at 4° C (SS 34 rotor of a Sorvall RC2-B centrifuge) at 31,000 x g for 10 min
to remove insoluble material. A fat cap was carefully removed from the surface with a
small spatula and supernatant fluid was decanted and filtered through cheesecloth to
remove particulate matter that was not pelleted. Cheesecloth was pre-soaked in

homogenization buffer to minimize loss of sample during the filtration process. The

20

supernatant fluid was then centrifuged at 4° C (rotor 30 of a Spinco model L ultra
centrifuge) at 85,000 x g for 30 minutes to remove the membrane fraction. The resulting
supernatant fluid obtained was adjusted to contain 5 mM ethylenediaminetetraaeetic acid
(EDTA) and 0.1 mM dithiothreitol. Then 7 mL of supernatant were placed on a column
(Bio-Rad Column 2.5 cm ID x 14 cm length, cat. No. 738-0017) packed with Sephadex
G-25 resin (fine; Amersham Pharmacia). Sephadex beads were prepared by soaking in
homogenization buffer. They were stirred every h for 3 h, and allowed to swell for a total
of 12 h prior to being transferred to the column. The column was flushed with 0.5 L of
homogenization buffer to ensure that all Sephadex beads were properly packed and
equilibrated prior to sample loading. Protein was eluted from the column with a buffer
containing 15 mM potassium phosphate, pH 6.5, 280 mM potassium chloride, 5 mM
EDTA, and 0.1 mM dithiothreitol at a rate of 2 mL per minute and collected in 0.5 mL
fractions. A Bio-Rad Biologic LP Chromatography System (Cat. No. 731-8300) and
Bio-Rad Model 2128 Fraction Collector were used for gel filtration and collection of
proteins. Protein concentrations of gel-filtered samples were determined using a
commercial kit (BioRad Protein Assay Kit, BioRad Laboratories, Hercules, CA) based on
the method of Bradford (1976). Bovine serum albumin was used as a protein standard.
Samples with the highest protein concentrations were pooled and frozen in 1 mL aliquots
and stored at -80° C for later use. Care was taken to remove all filtrate from the column
by flushing the system with an additional 0.5 L of elution buffer, as well as 0.5 L of de-
ionized distilled H20. The column was prepared for the next sample filtration by

flushing with 0.5 L of homogenization buffer.

21

P l amide Gel Ele oresis AGE Anal sis of Protein

Separating gels were used to visualize gross differences that may exist in protein
profiles of gel-filtered muscle extracts from superior and inferior quality pork samples
used in this study. Twenty-eight ug of muscle protein were loaded on discontinuous
12.5% SDS-PAGE separating gels (Laemmli, 1970; 12.5% acrylamide/bis [37.5:1 ratio
of acrylamide to N, N’-methylene—bis-acrylamide]; approximately 9.5 cm x 6 cm x 0.75
m) using a Bio-Rad Mini Protean II apparatus (Bio-Rad, Hercules, CA). Gels were
allowed to polymerize for 1 hour before a 4% stacker (4% acrylamide/bis [37.5:1 ratio of
acrylamide to N, N’- methylene-bis-acrylamideD was poured. The composition of the
running buffer was 25 mM Tris, 192 mM glycine, and 0.1% SDS, pH 8.3. Gels were run
at 200 volts at room temperature for 45 min or until the dye front ran off. After
electrophoretic separation, proteins were stained in Coomassie blue R-250 (40%
methanol, 7% acetic acid, and 0.0025% coomassie blue) overnight and gels were
destained (40% methanol, 10% acetic acid) prior to being dried using the GelAir
Cellophane (Bio-Rad cat. No.165-1779) and scanned using a Bio-Rad fluor-s image
analysis system (Bio-Rad, Hercules, CA).
Myofibril Boom

Crude myofibrils were prepared using a protocol modified from Lonergan et a1.
(1995). Porcine longissimus muscle tissue (1 g) obtained at 24 hours postmortem was
homogenized in 9 volumes of an extraction buffer containing 10 mM tris hydrochloride,
pH 8.3, 10 mM EDTA, and 2mM phenylrnethylsulfonylfluoride (PMSF). The sample
was then centrifuged at 27,000 x g for 30 min (SS 34 rotor of a Sorvall RC2-B

centrifuge). Supernatant fluid was decanted and the pellet was saved. The pellet was

22

then re-homogenized in 5.0 mL of extraction buffer and centrifuged at 1,500 x g for 10
min. This procedure was repeated three times to further rinse the myofibrils. Following
the third centrifugation, the supematant fluid was decanted and the pellet was re-
suspended in 5 volumes of 50% glycerol and stored in 200 uL aliquots at -80° C for later
use.

Aliquots (200 uL) were thawed and centrifuged at 1,500 x g for 10 minutes to
pellet myofibrils, and glycerol was discarded. Samples were subsequently re-suspended
in 50 mM triethanolamine, pH 7.5 prior to use in reaction mixture.

Glycolytic Regction Conms

Gel-filtered samples containing muscle protein were exposed to a reaction
mixture to initiate glycolysis as described by Tomheim et al. (1991) with minor
modifications. The reaction mixture was made in a 10X stock (20 mL), and the final
reaction was performed in a volume of 2 mL in a polypropylene rnicrocentrifuge tube that
contained 4 mM aspartate, 10 mM glucose, 7.5 mM orthophosphate, 25 mM imidazole
HCl, pH 6.9, 8.3 mM magnesium chloride, 0.2 mM calcium chloride, and 1 mM ATP,
0.3 mM GTP, and 0.1 mM NAD, 0.25 mg/mL crude myofibrils, and 1 mg/ml gel-filtered
protein. The reaction was started by adding 33.6 uL of a stock yeast hexokinase (5.0 pl
of hexokinase in 495 uL of de-ionized distilled H20, Sigma cat. No. H-5625) to the 2 mL
volume, which provided 0.06 units/mL hexokinase activity in the reaction. Tubes
containing reaction mixture were then inverted 3 times to mix the contents. The reaction
mixture was maintained in a water bath at 30° C. A portable pH meter with a puncture
type probe was used to measure pH at 0, 10, 20, 30, and 40 min following the initiation of

the reaction. Samples of 75 ul were removed every 2 minutes fiom 0 to 46 minutes and

23

placed in 75 uL of 2 N perchloric acid to deproteinize the sample. Upon completion of
the experiment, acidified samples were centrifuged at 1,500 x g for 10 minutes in an
Eppendorf 5415 centrifuge. Following centrifugation, perchloric acid in samples was
neutralized with 30 uL of 5.4 N KOH. The samples were then re-centrifuged at 1,500 x g
for 10 minutes. Supernatant fluid was kept on ice and assays were performed within 2
hours. Unless otherwise indicated, data represent results of an individual experiment.
ATP Assay

Adenosine triphosphate (ATP) was quantified using the enzymatic method of
Lamprecht and Trautschold (1972), with the exception that the assay was adapted to a 96-
well microtiter plate. The ATP assay reagent consisted of 50 mM triethanolarnine buffer,
10 mM NAD, 0.1 M magnesium chloride, and 0.5 M glucose. The total reaction volume
was 230 uL and the quantity of ATP was determined by following the conversion of
NAD+ to NADH at 340 nm using a VERSAmax plate reader (Molecular Devices,
Sunnyvale, CA). The coupled enzyme reactions that are the basis of this assay are shown

below.

ATP + Glucose i, G-6-P + ADP

l
I

G-6-P + NAD fl. 6-Phosphoglucono-lactone + NADH + HI
Assays were performed in duplicate with 10 uL of sample from the deproteinized

reaction mixture followed by 200 uL of assay reagent. An initial absorbance (A1) at 340

nm was recorded. A stock of glucose-6-phosphate dehydrogenase was prepared in H20,

24

and 10 uL(0.46 U/mLinfinalreactionmixture)wasaddedto eachwelltostartthe
reaction. The contents of each well were mixed and incubated at 37°C for 15 minutes,
andasecondAmreading(A2)wastaken. Thiswasamodificationofthe original
protocol. It was necessary to run the initial reaction to completion to avoid confounding
effects of residual G-6-P produced in the glycolytic reaction. This allowed for the
quantification of G-6«P (AZ-A1). A stock of hexokinase (Sigma cat. No. H-4502) was
also prepared and 10 uL (1.8 U/mL in final reaction mixture) was added to each well.
The contents of each well were mixed and incubated at 37°C for 40 minutes. Upon
completion of the reaction the absorbance at A340 (A3) was again recorded and the
change in absorbance (A3-A2) was used for ATP determination. Fresh ATP standards (1
mM, 0.5 mM, 0.25 mM, 0.125 mM, and 0 mM) were prepared daily, and were used to
create a linear regression equation (y = mx + b) that was implemented to calculate the
ATP concentration of sample unknowns. Final concentrations of ATP were multiplied
by a dilution factor of 2.4 to account for dilution of sample during acidification and
neutralization.
ADP Assay

The enzymatic determination of adenosine diphosphate (ADP) also involved the
utilization of methods from Lamprecht and Trautschold (1972). The assay involved an
initial creatine kinase reaction to quantitatively convert ADP to ATP. This reaction was
performed by adding 30 ul of a solution containing 14 mM creatine phosphate, 0.1 M
glycine, 1 mM magnesium sulphate, pH 9.0, and creatine kinase (25 U/mg), and 30 ul of
sample to microtubes. Reaction mixtures were incubated for 15 minutes at 37° C and

then heated for 3 min in a water bath at 95° C. Samples were subsequently centrifuged at

25

1,500 x g for 10 min. Then 20 uL ofthe sample were added to microtiter plate wells, and
the aforementioned ATP protocol was utilized to quantify ATP. The concentration of
ADP was determined as the difference between the ATP concentration measured before
and alter conversion of ADP to ATP using the creatine kinase reaction. Final
concentrations were derived as previously stated for ATP.
lactate Assay

lactate accumulation was measured in samples collected from the glycolytic
reaction procedure using methods fiom Sigma (Procedure No. 826—UV). The assay is

based on the reaction shown below.

Pyruvate + NADH ——.|”’" Lactate + NAD
The reaction was performed in a 96—well microtiter plate and the final reaction
volume was 210 "L. Samples (10 uL) were added in duplicate to each well followed by
200 uL of assay reagent which included 2.47 mM NAD, 0.6 mM glycine, and lactate
dehydrogenase (1,000 U/mL). Additionally, elution buffer (10 uL) and assay reagent
(200 uL) were used as a blank for this assay. Samples were then mixed and placed in an

incubator for 15 min at 37° C.

26

Upon completion of the reaction, lactate concentration was determined by
following the decrease in absorbance at 340 nm due to the oxidation of NADH to NADI.
Absorbance was measured on a VERSAmax plate reader (Molecular Devices, Stmnyvale,
CA), and final concentrations of lactate were calculated using the following equation,

where 6.22 represents millimolar absorptivity of NADH at 340 nm.

AA34o "' Total Volume = AA340 ° 210 uL
6.22 mL/umol/cm “ Sample Vol. * Light Path 6.22 mL/umol/cm * 10 uL "' 0.65 cm

 

27

RESULTS AND DISCUSSION

An in vitro glycolytic system was used to determine if muscle protein extracts
fiom pigs that produced superior and inferior quality pork (Table 1) exhibited different
patterns of glycolysis. Loins from pigs designated superior quality had higher 45 min
pH, lower fluid loss in samples subjected to centrifugal force, lower drip loss, and lower
Day 1 L" values than loins fiom pigs designated inferior in quality (Table 1).

Supernatant from longissimus muscle extracts was gel-filtered and the collected
fiactions (F igure 1) were pooled from pigs representing both types of meat quality. Then
fractions were analyzed on an SDS PAGE gel to determine molecular weights and
similarities of protein bands (Figure 2, lanes 2 to 7). No apparent differences exist in the
protein profiles for the superior and inferior quality samples. In addition to sarcoplasmic
proteins (~16.5 to 97 kDa), pig muscle extracts also contain relatively abundant myosin
(~200 kDa). The cause for the prominent myosin band could be the relatively high ionic
strength of the homogenization buffer used compared to buffers typically used for
extraction of sarcoplasmic proteins or myofibrils (Pearson and Young, 1989). The fact
that samples were harvested from pre-rigor loin muscle may also contribute to the
abundance of myosin in the extract, since myosin is soluble at an ionic strength of 0.23
whereas actomyosin is not (Pearson and Young, 1989). Contrary to the pig muscle,
myosin is less evident in the rat hind limb extract (Figure 2, lane 8). This is most likely
attributable to the rat muscle tissue being closer to the completion of rigor than the

porcine muscle used in this study.

28

Postmofl Engrgy Utilization
Oscillatory behavior of glycolysis in skeletal muscle extracts is caused by

repeated bursts of PFK activity and associated oscillations in the ATP/ADP ratio. This
involves the AMP-dependent activation of PFK by its product, fructose-1 ,6 diphosphate.
When the ATP/ADP ratio lowers to a trigger level (ratio of approximately 2 to 4), this
initiates a sudden burst in glycolytic flux that restores a high ATP/ADP ratio (T omheim
et al., 1991). This pattern of glycolytic behavior was observed by Tomheim et al., (1988,
1991) in gel-filtered rat hind limb muscle extracts.

We anticipated that pigs with superior quality traits would exhibit more subtle
glycolytic oscillations, while pigs with inferior quality traits would exhibit more
pronounced oscillatory patterns of energy production. However, when muscle protein
fractions were exposed to glycolytic reaction mixtures, a variety of patterns of glycolysis
were observed.

The largest fluctuation in adenine nucleotide concentration occurred from 0 to 20
min following the initiation of the glycolytic reaction in samples of most pigs. Thus,
samples were classified as either oscillatory or non-oscillatory based on their pattern of
energy generation during the first 20 min. Figures 3.1, 3.2, and 3.3, respectively, depict
non-oscillatory patterns of glycolysis. The rate of glycolysis in these samples appears to
parallel the rate of ATP utilization, such that the ATP concentration deviates little, while
lactate steadily accumulates. This pattern of glycolysis was hypothesized to be
associated with superior quality pigs, but was exhibited by both superior and inferior
quality pigs. In contrast, figures 3.4, 3.5, and 3.6, respectively, depict an oscillatory

pattern of glycolysis. This oscillatory pattern was hypothesized to occur in inferior

29

quality pigs and be associated with localized acidification that may exacerbate protein
denaturation and increase fluid loss from product. However, the oscillatory pattern was
observed in both inferior and superior quality pigs.

As expected, ATP and ADP concentrations are inversely related to each other in
most samples, regardless of quality designation. There were no differences in the
average ATP/ADP ratio or the overall mean concentrations of lactate (Figure 4) or
adenine nucleotides in reactions using extracts from either superior or inferior quality loin
samples (Table 2). Additionally, sire breed had no effect on the concentrations of
adenine nucleotides or lactate. Likewise, no differences were observed in the in vitro pH
decline between the superior or inferior quality samples (Figure 5). This is in contrast to
the more rapid pH decline that resulted in lower 45 min pH of inferior quality loins
(Table 1). Although conditions such as temperature and calcium concentration were
controlled in the in vitro assay, variation in these traits may influence the rate of
postmortem glycolysis in carcass muscles. Nonetheless, our data suggest that protein
extracts fiom muscle yielding superior and inferior quality pork do not produce distinct
patterns of glycolysis that would explain differences in pork loin quality. Regulation of
the glycolytic pattern may be attributed to differences in metabolic effectors or the rate of
energy utilization during postmortem glycolysis.

We would expect more pronounced oscillations that occurred from 0 to 20 min to
be accompanied by greater H+ ion accumulation in a localized environment, such as in
proximity to the calcium ATPase, where glycolytic enzymes have been shown to be
functionally compartmentalized (Xu and Becker, 1998). Ifthis occurred in muscle, one

would expect to incur greater changes in color attributes and WHC due to protein

30

denaturation. However, analysis of the glycolytic pattern throughout the entire time
period (0 to 46 min) may be of significance when making implications related to pork
quality. All samples exhibited an oscillatory pattern (2 2 oscillations) of glycolysis fiom
20 to 46 min, but the amplitude of these oscillations was lower than those observed from
0 to 20 min. It would appear that reduced amplitude of oscillations occurring fi'om 20 to
46 min may be a result of the system losing finite control over glycolysis due to low pH.
At the onset of the reaction, pH is near neutral and substrate is plentiful. As the reaction
proceeds and H” ions accumulate, the pH is driven downward to ~ 5.7 (Figure 5). As pH
continues to fall, the rate of glycolysis slows, ATP production decreases, and lactate
accumulates at a reduced rate. Tomheim and Lowenstein (1973) have previously shown
that glycolytic oscillations are strongly dependent upon pH. These authors observed that
at lower pH, deamination occurred to a greater extent and more time elapsed before
reamination.

When compared to rat hind limb muscle extract (Tomheim et al., 1979), pig
muscle extract resulted in more rapid in vitro glycolysis. A comparison of glycolytic
behavior between pig and rat muscle extracts would determine if species or laboratory
techniques were responsible for the deviation in energy production. Figure 6 depicts the
oscillatory pattern of glycolysis that we achieved using rat hind limb muscle extract.
There appears to be a decrease in the rate and extent of lactate accumulation in the rat
hind limb muscle when compared to the protein extracts from pig muscle. Our pattern of
ocillatory behavior for rat was similar to Tomheim et al. (1974, 1979) where the rate of
reaction in extracts from rat hind limb muscle was much slower. Despite the slower rate

of lactate accumulation, deamination of adenine nucleotides was evident in reactions

31

containing rat muscle extract. This was shown in the sharp decline in concentrations of
ATP and ADP seen hour 20 to 46 min during the reaction period. The decreased lactate
accumulation in the rat muscle extract is likely to be attributable to fiber type differences,
as rat hind limb would be expected to have a higher pr0portion of red fibers, less
glycolytic activity, and lower ATPase activity than pig longissimus muscle (Pearson and
Young, 1989).
Adding an Alternative ATPase

It is well established that the rate of ATP utilization is positively associated with
the rate of pH decline. For example, species with a higher proportion of white muscle
fibers, such as pigs and poultry, undergo more rapid postmortem ATP utilization and
glycolysis than red muscle species (i.e. beef and lamb) due to more active myosin
ATPase and more abundant SR calcium ATPase. Increased ATPase activity may reduce
the ATP/ADP ratio and induce production of glycolytic “burst” activity. To determine
the effect of added ATPase from a myofibrillar source on the pattern of glycolysis,
protein extracts from a pig of each quality type (1 mg/mL of gel-filtered protein) were
used in combination with 0, 0.25, and 0.50 mg/mL of myofibrils. Although the added
ATPase caused increased lactate accumulation over the reaction period of 0 to 46 min,
the pattern of glycolysis was not affected by the addition of myofibrils (Fig. 7.1 and 7.2).

How (or if) the functional coupling of glycolytic enzymes to the major sites of
energy utilization (myosin ATPase and calcium ATPase) affect pork quality is currently
unknown. However, the effects of this coupling may influence glycolytic rate, ultimate
enzyme location and the degree of denaturation of sarcoplasmic proteins, which, in turn,

may influence the color (Joo et al., 1999) and WHC of pork (Wilson and van Laack,

32

1999). The functional coupling of glycolytic enzymes to the myofibrillar ATPase was
beyond the scope of these experiments.
Effects of Citrate

The experiments previously described demonstrated that pig muscle extracts
produce glycolysis with an oscillatory behavior, but no difl‘erences were observed in the
visual pattern of glycolysis using extracts from superior and inferior pork. Furthermore,
no differences were observed in mean concentration of adenine nucleotides or lactate
over all time points (P > 0.05). Thus, if patterns of glycolysis differ between postmortem
muscles that exhibit variation in meat quality, these patterns are not mimicked by extracts
containing glycolytic enzymes from those muscles.

Although glycolytic enzyme capacity does not appear to explain a large
proportion of the variation in pork color and WHC, other levels of glycolytic regulation
may contribute to the rapid pH decline often associated with PSE pork. Citrate is known
to be a potent physiological inhibitor of PFK, which is generally regarded as the primary
regulatory enzyme of glycolysis. Tomheim et al. (1991) showed that addition of citrate
in extracts of rat hind limb muscle decreased the frequency of the oscillations and
delayed the first burst of PFK activity. These effects are due to inhibition of PFK, such
that a lower trigger value of the [ATP]/[ADP] ratio must be reached before a burst of
PFK can be initiated. When 0.15 mM citrate was included in the reaction mixture
containing pig muscle extract, the rate of lactate accumulation was decreased (Figure 8).
Our results support Tomheim’s previous findings, and demonstrate that the in vitro

system used is responsive to known physiological regulators of glycolysis.

33

Similar to citrate, fructose 2,6-bisphosphate has a regulatory role in glycolytic
behavior. Fructose 2,6-bisphosphate is a potent activator of PFK and is competitive with
fructose 1,6-bisphosphate for the same binding site on PFK. Tomheim et al. (1988) have
shown that addition of fructose 2,6-bisphosphate will decrease the ATP/ADP ratio,
essentially blocking oscillatory patterns of glycolysis. This blockage is the result of
autocatalytic activation of fructose 1,6-bisphospate, and thus prevents glycolytic
oscillations. With this said, it is apparent that control over glycolytic flux, or the flow of
intermediates through glycolysis, as well as regulatory mechanisms influencing

postmortem metabolism warrant further research.

34

IMPLICATIONS

Today’s competitive agricultural economy has constricted the profit margins
experienced in the swine industry. Consequently, it’s not a surprise that costly problems
related to inferior meat quality warrant further research In this study, a system was
successfully developed to monitor in vitro glycolytic energy metabolism. Oscillatory
patterns of glycolysis in vitro do not appear to be related to deviations in pork quality
measured today. However, this system will permit identification of specific metabolites
that cause differences in the pattern of glycolysis. Further research is needed to monitor

the influence of individual metabolites on the behavior of glycolytic metabolism.

35

APPENDICES

Table 1. Means and standard deviations for carcass quality data”

 

_p_en__Qu_ahat ___QUAM'OI
Stdev Mean Stdev

45 min pH 6.40 0.14 5.92 0.24
CWHC,% 10.22 1.15 20.70 0.78
Drip 1,% 0.66 0.19 3.23, 1.10
Day 1 L‘ 51.37 1.61 56.68 2.11

 

' n = 6 pigs in each quality category (superior and inferior).

Table 2. Mean concentrations of adenine nucleotides and lactate observed in
glycolytic reactions from 0 to 46 min using protein extracts from either superior
or inferior quality pork. ‘b

 

Su rior i Inferior Qu_alig

Mean Stdev Mean Stdev
ATP, mM 0.539 0.256 0.610 0.268
ADP, mM 0.216 0.137 0.192 0.120
ATP/ADP 4.506 3.390 5.361 3.239
Lactate l .707 0.790 1 .673 0.745

 

 

'n= 6 pigs in each quality category (superior and inferior).
b No differences were observed 1n reactions using extracts fiom superior and
inferior quality samples (P > 0. 05).

36

Protein (mg/mL)

 

o 5 1o 15 20 25
Fraction Number
Figure 1. Profile of pig muscle protein eluted from gel filtration column. Fractions of
0.5 mL were collected. Protein eluted between the arrows was pooled and used in

reaction mixtures. Gel-filtration was used to remove small molecular weight substances
that potentially influence glycolysis.

37

C
M
D—§ “r -‘ m n—ro-fi ,. m“ al.-his
an: are: _ an.

 

F —> k
"~' ~~- “re-4w 1» muck w“...
3‘- ion”: ‘» News? '5... ya:
G—D \‘k
m. m er .. has.» ... ,2. a“
., g... ""'" .
H—> \_ ,- .2-. ~ -. "M
‘

Figure 2. Electrophoretic separation of proteins derived from gel filtration. A 12.5%
polyacrylarnide separating gel was used to search for crude deviations in the protein
profiles of muscle extracts differing in quality. Lane 1 includes molecular weight
markers. Lanes 2, 4, and 6 depict protein profiles of pigs designated superior quality.
Lanes 3, 5, and 7 depict protein profiles of pigs designated inferior quality. Lane 8
depicts the protein profile of rat hind limb muscle. Lane 9 depicts the protein profile of a
HAL-1843 positive pig. Twenty-eight 11g of muscle protein were loaded in each lane.
Rows A -— H represent differing molecular weight markers as follows: A = 200.0 kDa; B
= 116.3 kDa; C = 97.4 kDa; D = 66.2 kDa; E = 45.0 kDa; F = 31.0 kDa; G = 21.5 kDa; H
= 14.4 kDa.

38

Superior Quality - Duroc (316#8)

+Lactate
+A’1'P
+ADP

 

Figure 3.1. Adenine nucleotide (ADP and ATP) and lactate concentrations of a superior
quality Duroc pig exhibiting non-oscillatory glycolytic behavior from 0 to 46 min after
initiation of glycolytic reaction.

39

Superior Quality - Pietrain (322#5)

+Lactate
+A'rp

 

Figure 3.2. Adenine nucleotide (ADP and ATP) and lactate concentrations of a superior
quality Pietrain pig exhibiting non—oscillatory glycolytic behavior from 0 to 46 min after
initiation of glycolytic reaction.

Inferior Quality - Duroc (322#6)

+ ATP
+ADP

 

Time (min)

Figure 3.3. Adenine nucleotide (ADP and ATP) and lactate concentrations of an inferior
quality Duroc pig exhibiting a non-oscillatory pattern of glycolytic behavior from 0 to 46
min after initiation of glycolytic reaction.

41

Inferior Quality - Pietrain (326#7)

+1.30th
+ATP
+ADP

 

Figure 3.4. Adenine nucleotide (ADP and ATP) and lactate concentrations of an inferior
quality Pietrain pig exhibiting oscillatory glycolytic behavior from 0 to 46 min afier
initiation of glycolytic reaction.

42

Superior Quality - Pietrain (322#8)

+lactate
+ ATP
+ ADP

 

Time (min)

Figure 3.5. Adenine nucleotide (ADP and ATP) and lactate concentrations of a superior
quality Pietrain pig exhibiting an oscillatory pattern of glycolytic behavior from 0 to 46
min after initiation of glycolytic reaction.

43

Inferior Quality - Duroc (320#8)

+ lactate
+ ATP
+ ADP

 

Figure 3.6. Adenine nucleotide (ADP and ATP) and lactate concentrations of an inferior
quality Duroc pig exhibiting an oscillatory pattern of glycolytic behavior from 0 to 46
min after initiation of glycolytic reaction.

Superior vs Inferior Quality - lactate Accumulation (0 to 46min)

  
 

‘ 3 + Superior Quality
i + Inferior Quality

Figure 4. Lactate accumulation from 0 to 46 min after initiation of glycolytic reaction.
The data is indicative of the means for each time point of the respective quality groups
(superior vs inferior). N = 6 pigs in each quality category (superior and inferior).

45

pH Measures in Superior vs Inferior Muscle Extracts During Glycolytic
Reaction

+ Superior Quality
+ Inferior Quality

 

Figure 5. In vitro pH decline of superior and inferior quality samples during reactions (0
to 46 min) performed using the glycolytic conditions described in the materials and
methods. N = 6 pigs in each quality category (superior and inferior).

+lactate
+ATP
+ADP

 

Figure 6. Adenine nucleotide (ADP and ATP) and lactate concentrations of rat hind limb
muscle exhibiting an oscillatory pattern of glycolytic behavior flour 0 to 46 min afier
initiation of glycolytic reaction.

47

Superior Quality (316#8) - lactate Accumulation

+ 0 mg/mL Myofibrils
+ 0.25 mg/mL Myofibrils
+ 0.50 mg/mL Myofibrils

 

Time (min)

Figure 7.1. Lactate accumulation of a supeior quality Duroc pig with the addition of
three differing concentrations of myofibrils from 0 to 46 min alter initiation of glycolytic
reaction.

48

Inferior Quality (322#6) - Lactate Accumulation

+ 0 mg/mL Myofibrils
-l- 0.25 mg/mL Myofibrils
+ 0.50 mg/mL Myofibrils

 

0 10 20 30 4o 50
Time (min)
Figure 7.2. Lactate accumulation of an inferior quality Duroc pig with the addition of

three differing concentrations of myofibrils from 0 to 46 min after initiation of glycolytic
reaction.

49

Effects of Citrate on Lactate Accumulation

+ Citrate Added
-I— w/o Citrate

 

Figure 8. Lactate concentration of pig muscle extract exposed to differing concentrations
of citrate from 0 to 46 min after initiation of glycolytic reaction.

50

LITERATURE CITED

Allison, C.P., R.0. Bates, A.M. Booren, and ME. Doumit 2003. Pork quality variation
is not explained by glycolytic enzyme capacity. Meat Sci. 63:7-22.

Alonso, M.D., J. Lomako, and WJ. Whelan. 1995. A new look at biosynthesis of
glycogen. FASEB J. 9:1126-1137.

Bechtel, PJ. and P.M. Best 1985. Integration of ATP production and consumption.
Ree. Meat Conf. Proc. 38:26-31.

Bendall, J .R., and J. Winner-Pedersen. 1962. Some properties of the fibrillar proteins of
normal and watery pork muscle. J. Food Sci. 27:144-159.

Bertram, H.C., P.P. Purslow, and HJ. Andersen. 2002. Relationship between meat
structure, water mobility, and distribution: A low-field nuclear magnetic
resonance study. J. Agric. Food Chem. 50:824-829.

Boles, J .A., F .C. Parrish, Jr., T.W. Huiatt, and RM. Robson. 1992. Effect of porcine
stress syndrome on the solubility and degradation of myofibrillar/cytoskeletal
proteins. J. Anim. Sci. 70:454-464.

Boles, J .A., J.F. Patience, A.L. Schaefer, and IL. Aalhus. 1994. Effect of oral loading of
acid base on the incidence of pale soft exudative pork (PSE) in stress susceptible
pigs. Meat Sci. 37:281-194.

Briskey, El 1964. Etilogical status and associated studies of pale, soft, exudative
porcine musculature. Adv. Food Res. 13:89-105.

Briskey, E.J., R.W. Bray, W.G. Hoekstra, P.H. Phillips and RH. Grummer. 1959. The
chemical and physical characteristics of various pork ham muscle classes. J.
Anim. Sci. 18:146.

Briskey, E.J., L.L. Kastenschmidt, J .C. Fonest, G.R. Beecher, M.D. Judge, R.G. Cassens,
and W.G. Hoekstra. 1966. Biochemical aspects of post-mortem changes in
porcine muscle. J. Agric. Food Chem. 14:201-207.

Briskey, EJ. and J. Wismer-Pedersen. 1961. Biochemistry of pork muscle structure. 11.
Preliminary observations of biopsy samples versus ultimate muscle structure. J.
Food Sci. 26:306-313.

Cameron, ND. 1990. Genetic and phenotypic parameters for carcass traits, meat and
eating quality traits in pigs. Livest. Prod. Sci. 26:119-135.

51

Cannon, J .E., J.B. Morgan, J. Heavner, F .K. McKeith, G.C. Smith and D.L. Meeker.
1995. Pork quality audit: A review of the factors influencing pork quality. J.
Muse. Foods. 6:369-402.

Cannon, J.E., J .8. Morgan, J. Heavner, F .K. McKeith, G.C. Smith, and D.L. Meeker.
1996. Pork quality audit survey: Quantification of pork quality characteristics. J.
Muse. Foods. 7:29-44.

Cassens, R.G. 2000. Historical perspectives and current aspects of pork meat quality in
the USA. Food Chem. 69:357-363.

Cheah, K.S., and A.M. Cheah. 1976. The trigger for PSE condition in stress-susceptible
pigs. J. Sci. Food Agric. 27:1137-1144.

Ciobanu, D., J. Bastiaansen, M. Malek, J. Helm, J. Woollard, G. Plastow, and M.
Rothschild. 2001 . Evidence for new alleles in the protein kinase adenosine
monophosphate activated 73-subunit gene associated with low glycogen content in
pig skeletal muscle and improved meat quality. Genetics. 159:1151-1162.

Conley, K.E., M.L. Blei, T.L. Richards, MJ. Kushmerick, and SA. Jubrias. 1997.
Activation of glycolysis in human muscle in vivo. Am. J. Physiol. 273:C306-
C315.

Connett, RJ., and K. Sahlin. Control of Glycolysis and glycogen metabolism.
Handbook of Physiology, Oxford University Press, NY. 1996. P870-911.

Deng, Y., K. Rosenvold, A.H. Karlsson, P. Horn, J. Hedegaard, C.L. Steffensen, and H.J.
Andersen. 2002. Relationship between thermal denaturation of porcine muscle
proteins and water-holding capacity. J. Food Sci. 67:1642-1647.

Doumit, M.E., C.P. Allison, and M.J. Ritter. 2001. Control of glycolysis in relation to
pork quality traits. Reciprocal Meat Conference Proceedings. 54:155-157.

Dutson, TR. 1983. The measurement of pH in muscle and its importance to meat
quality. Reciprocal Meat Conference Proceedings. 36:92-97.

Fujii, J ., K. Ostu, F. Zorzato, S.D. Leon, V.K. Khama, J.B. Weiler, P.J. O’Brien, and
DH. MacLennan. 1991. Identification of a mutation in porcine ryanodine
receptor associated with malignant hyperthermia. Science. 253:448-451.

Garrett, RH. and CM. Grisham. 1999. Chapter 17: Molecular motors in biochemistry.
2"‘1 ed. p 533-564. Saunders College Publishing, Orlando, FL.

Gerrard, DE. 1997. Pork quality: Beyond the stress gene. National Swine Improvement
Federation Proceedings. Vol. 22.

52

6011, DE, R.M. Robson, and M.H. Stromer. 1984. Skeletal Muscle. In:M.J. Swenson
(ed) Duke’s physiology of Domestic Animals. 10'“ ed. p 548-580. Cornell
University Press, Ithaca, NY.

G011, D.E., V.F. Thompson, R.G. Taylor, and J.A. Christiansen. 1992. Properties and
regulation of the calpain system and its role in muscle growth. Biochimie.
74:225-237.

Honikel, K0. and OJ. Kim. 1986. Causes of the development of PSE pork.
Fleischwirtsch. 66:349-353.

Honikel, K0. and R. Hamm. 1994. Advances in meat research, Vol. 9 (eds. A.M.
Pearson and TR. Dutson). Chapman and Hall, Glasgow, UK.

Huang, M., C. Lee, R. Lin, and R. Chen. 1997. The exchange between proglycogen and
macroglycogen and the metabolic role of the protein rich glycogen in rat skeletal
muscle. J. Clin. Invest. 99:501-505.

Huff-Lonergan, E., F .C. Parrish, Jr., and R.M. Robson. 1995. Effects of postmortem
aging time, animal age, and sex on degradation of titin and nebulin in bovine
longissimus muscle. J. Anim. Sci. 73:1064-1073.

Huff-Lonergan, E., T. Mitsuhashi, D.D. Beekman, F .C. Parrish, D.G. Olson and R.M.
Robson. 1996. Proteolysis of specific muscle structural proteins by u—calpain at
low pH and temperature similar to degradation in postmortem bovine muscle. J.
Anim. Sci. 74:993-1008.

lvers, D.J., L.F. Richardson, D.J. Jones, L.E. Watkins, K.D. Miller, J.R. Wagner, R.
Seneriz, A.G. Zimmennann, K.A. Bowers, and DB. Anderson. 2002.
Physiological comparison of downer and non-downer pigs following
transportation and unloading at a packing plant. Midwest Animal Science
Meetings Proceedings. 36:10.

Joo, S.T., R.G. Kauffrnan, B.C. Kim, and GB. Park. 1999. The relationship of
sarcoplasmic and myofibrillar protein solubility to colour and water-holding
capacity in porcine longissimus muscle. Meat Sci. 52:291-297.

Kastenschmidt, L.L., W.G. Hoekstra, and El Briskey. 1968. Glycolytic intermediates
and co-factors in “fast-” and “slow-glycolyzing” muscles of the pig. J. Food Sci.
3 3: 15 1-1 58.

Kauffman, R.G., D. Waccholz, D. Henderson and J .V. Lochner. 1978. Shrinkage of

PSE, normal and DF D hams during transit and processing. J. Anim. Sci.
46:1236-1240.

53

Kauffman, R.G., R.G. Cassens, A. Scherer, and D.L. Meeker. 1992. Variations in pork
quality: history, definition, extent, resolution. National Pork Producers Council
Publication. Des Moines, Iowa.

Kauffman, R.G., R.L. Russell, and M.L. Greaser. 1999. Using pork to teach students
quality variations and how they are measured. J. Anim. Sci. 77:2574z2577.

Kauffman, R.G., S.T. Joo, C. Schultz, R Warner, and C. Faustman. 1994. Measuring
water-holding capacity in post-rigor muscle. Reciprocal Meat Conference
Proceedings. 47:70-72.

Koohmaraie, M. 1992. The role of Cay-dependent protease (calpains) in postmortem
proteolysis and meat tenderness. Biochimie. 74:239-245.

Koohmaraie, M. 1996. Biochemical factors regulating the toughening and tenderization
process of meat. Meat Sci. 43:Sl93-S201.

Koohmaraie, M., G. Whipple, D.H. Kretchmar, J .D. Crouse and H.J. Mersmann. 1991.
Postmortem proteolysis in longissimus muscle from beef, lamb and pork
carcasses. J. Anim. Sci. 69:617-624.

Krause, U. and G. Wegener. 1996. Control of glycolysis in vertebrate skeletal muscle
during exercise. Am. J. Physiol. 270:R821-R829.

Laemmli, UK. 1970. Cleavage of structural proteins during the assembly of the head of
bacteriophage T4. Nature (Lond.) 227:680-685.

Lee, J.C., L.K. Hesterberg, M.A. Luther, and G-Z. Cai. Allosteric Enzymes. G. Herve
(Ed.). CRC Press, Florida. 1989. p 231-254.

Le Roy, R, J .M. Elsen, J .C. Caritez, A. Talmant, H. Juin, P. Sellier, and G. Monin. 2000.
Comparison between the three porcine RN genotypes for growth, carcass
composition and meat quality traits. Genet. Sel. Evol. 32:165-186.

Lomako, J ., M. Lomako, W.J. Whelan, R.S. Dombro, J .T. Neary, and MD. Norenberg.
1993. Glycogen synthesis in the astrocyte: from glyogenin to proglycogen to
glycogen. FASEB J. 7:1386-1393.

Lonergan, S.M., E. Huff-Lonergan, L.J. Rowe, D.L. Kuhlers, and SB. Jungst. 2001.
Selection for lean growth efficiency in Duroc pigs influences pork quality. J.
Anim. Sci. 79:2075-2085.

Louis, C.F., W.E. Rempel, and JR. Mickelson. 1993. Porcine stress syndrome:

Biochemical and genetic basis of this inherited syndrome of skeletal muscle.
Reciprocal Meat Conference Proceedings. 46:89-96.

54

Milan, 0., J .T. Jean, C. Loofi, V. Amarger, A. Robic, M. Thelander, C. Rogel-Gaillard,
S. Paul, N. Iannuccelli, L. Rask, H. Ronne, K. Lundstrtim, N. Reinsch, J. Gellin,
E. Kalm, P. Le Roy, P. Chardon, and L. Andersson. 2000. A mutation in
PRKAG3 associated with excess glycogen content in pig skeletal muscle.
Science. 288:1248-1251.

Monin, G. and P. Sellier. 1985. Pork of low technological quality with a normal rate of
muscle pH fall in the immediate post mortem period: The case of the Hampshire
breed. Meat Sci. 13:49-63.

Murray, A.C. and C.P. Johnson. 1998. Impact of the halothane gene on muscle quality
and pre-slaughter deaths in Western Canadian pigs. Can. J. Anim. Sci. 78:543-
548.

National Pork Producers Council (NPPC). 1991. Procedures to evaluate market hogs.
Des Moines, Iowa. 3rd ed.

National Pork Producers Council (NPPC). 1998. Pork quality targets. Des Moines,

Iowa. vmwvnppcorg.

 

National Pork Producers Council (NPPC). 2000. Pork composition and quality
assessment procedures. Des Moines, Iowa

National Pork Producers Council (NPPC). 2003. Pork facts. Des Moines, Iowa.

\untnppeorg.

 

Offer, G. 1991. Modeling of the formation of pale, soft and exudative meat: Effect of
chilling regime and rate and extent of glycolysis. Meat Sci. 30:157-184.

Parkhouse, W.S. 1992. Regulation of skeletal muscle metabolism by enzyme binding.
Can. J. Physiol. Pharrnacol. 70:150-156.

Pate, E. and R. Cooke. 1989. A model of crossbridge action: The effect of ATP, ADP
and P;. J. Muscle Res. Cell Motil. 10:181-196.

Pearson, A.M. and RB. Young. Muscle and meat biochemistry. Academic Press, Inc.,
San Diego, CA. 1989. Pl-457.

Purslow, P.P., A. Shafer, L. Kristensen, H.C. Bertram, K. Rosenvold, P.R. Henckel, H.J.
Andersen, P.J. Knight, T.J. Wess, S. Stoier, and M. Aaslyng. 2001. Water-
holding of pork: understanding the mechanisms. Reciprocal Meat Conference
Proceedings. 54:134-142.

Sayre, RN. and E.J. Briskey. 1963. Protein solubility as influenced by physiological
conditions in the muscle. J. Food Sci. 28:675-679.

55

Sayre, R.N., E.J. Briskey, and W.G. Hoekstra. 1963. Comparison of muscle
characteristics and post-mortem glycolysis in three breeds of swine. J. Anim. Sci.
22:1012-1020.

Schaefer, A.L., P.L. Dubeski, J.L. Aalhus, and AK. Tong. 2001. Role of nutrition in
reducing antemortem stress and meat quality abenations. J. Anim. Sci. 79:E91-
E101.

Schwagele, F ., C. Haschke, K.0. Honikel, and G. Krauss. 1996. Enzymological
investigations on the causes for the PSE-syndrome. Comparative studies on
pyruvate kinase from PSE- and normal pig muscles. Meat Sci. 44:27-39.

Scopes, RK 1964. The influence of post-mortem conditions on the solubilities of
muscle proteins. Biochem. J. 91 :201-207.

Scopes, R.K. 1977. Purification of glycolytic enzymes by using affrnity-elution
chromatography. Biochem. J. 161:253-263.

Sc0pes, R.K. and RA. Lawrie. 1963. Postmortem [ability of skeletal muscle proteins.
Nature 197:1202.

Scopes, R.K. 1974. The rate and extent of glycolysis in stimulated post-mortem
conditions. Biochem. J. 142279-86.

Smolen, P. 1995. A model for glycolytic oscillations based on skeletal muscle
phosphofructokinase kinetics. J .Theor. Biol. 174:137-148.

Stetzer, A.J., F.K. McKeith. Quantitative strategies and opportunities to improve quality.
Final Report, Benchmarking Value in the Pork Supply Chain. AMSA, Savoy, IL.
2003. P1-6.

Suarez, R.K. 1996. Upper limits to mass-specific metabolic rates. Annu. Rev. Physiol.
58:583-605.

Termonia, Y. and J. Ross. 1981. Oscillations and control features in glycolysis:
Numerical analysis of a comprehensive model. Proc. Natl. Acad. Sci. USA.
78:2952-2956.

Thomas, S. and DA. Fell. 1998. A control analysis exploration of the role of ATP
utilization in glycolytic-flux control and glycolytic-metabolite-concentration
regulation. Eur. J. Biochem. 258:956-967.

Tomheim, K. 1988. Fructose 2,6-bisphosphate and glycolytic oscillations in skeletal
muscle extracts. J. Biol. Chem. 263 :2619-2624.

56

Tomheim, K. and J.M. Lowenstein. 1973. Oscillations in metabolite concentrations
during the operation of the cycle in muscle extracts. J. Biol. Chem. 248:2670—
2677.

Tomheim, K. and J .M. Lowenstein. 1974. Interactions with oscillations of the glycolytic
pathway in muscle extracts. J. Biol. Chem. 249:3241-3247.

Tomheim, K., V. Andre's, and V. Schultz. 1991. Modulation by citrate of glycolytic
oscillations in skeletal muscle extracts. J. Biol. Chem. 266(24):15675-15678.

Tomheim, K. 1979. Oscillations of the glycolytic pathway and the purine nucleotide
cycle. J. Theor. Biol. 79:491-541.

United States Department of Agriculture (USDA). 2002. Agricultural statistics.
www.usda.gov/nass/pubs/agstats.htm.

Van Laack, R.L.J.M., S.G. Stevens and K.J. Stalder. 2001. The influence of ultimate pH
and intramuscular fat content on pork tenderness and tenderization. J. Anim. Sci.
79:392-397.

Warner, R.D., R.G. Kauffman, and M.L. Greaser. 1997. Muscle protein changes post
mortem in relation to pork quality traits. Meat Sci. 45:339-352.

Warriss, PD. and SN. Brown. 1987. The relationship between initial pH, reflectance
and exudation in pig muscle. Meat Sci. 20:65-74.

Warriss, P.D., E.A. Bevis, and P.J. Ekins. 1989. The relationships between glycogen
stores and muscle ultimate pH in commercially slaughtered pigs. Br. VetsJ.
145:378-383.

Wheeler, T.L., S.D. Shackelford, and M. Koohmaraie. 2000. Variation in proteolysis,
sacromere length, collagen content and tenderness among major pork muscles. J.
Anim. Sci. 78:958-965.

Wilson, G.G., III and KL. van Laack. 1999. Sarcoplasrnic proteins influence water-
holding capacity of pork myofibrils. J. Sci. Food Agric. 79:1939-1942.

Wismer-Pedersen, J. and E.J. Briskey. 1961. Rate of anaerobic glycolysis versus
structure in pork muscle. Nature 189:318.

Xu, K.Y., J.L. Zweier, J .L., and LC. Becker. 1995. Functional coupling between
glycolysis and sarcoplasmic reticulum Ca2+ transport. Circ. Res. 77:88-97.

Xu, K.Y., L.C. Becker. 1998. Ultrastructural localization of glycolytic enzymes on

sarcoplasmic reticulum vesicles. J. Histochernistry and Cytochemistry. 46:419-
427.

57

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