THE RELATIONSHIP BETWEEN ONION THRIPS (THRIPS TABACI) AND THE
BACTERIAL PATHOGENS PANTOEA AGGLOMERANS AND PANTOEA ANANATIS
By
Ari S. Grode

A THESIS
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
Entomology – Masters of Science
2017

ABSTRACT
THE RELATIONSHIP BETWEEN ONION THRIPS (THRIPS TABACI) AND THE
BACTERIAL PATHOGENS PANTOEA AGGLOMERANS AND PANTOEA ANANATIS
By
Ari S. Grode
Onion thrips (Thrips tabaci), a cosmopolitan pest of onion, are known vectors of Pantoea
agglomerans, causal agent of bacterial leaf blight, and P. ananatis, causal agent of bacterial
center rot. The objectives of this thesis were to determine the relationship between onion thrips
and P. agglomerans under field conditions, investigate the effectiveness of combining
insecticides and bactericides for managing both onion thrips and bacterial leaf blight and
determine the role of feeding injury by onion thrips in influencing the pathogenicity of P.
ananatis. In the first study, positive correlations between thrips population density, proportion of
thrips positive for P. agglomerans and bacterial leaf blight incidence in onion fields were
determined. In the pesticide trial conducted in 2015 and 2016, insecticides reduced both onion
thrips abundance and bacterial leaf blight incidence, while bactericides showed no effect on
reducing bacterial leaf blight symptoms. In the laboratory study significant positive correlations
were determined between the number of thrips per plant and the severity of leaf blight symptoms
as well as the percentage of feeding damage and the severity of leaf blight symptoms. Lastly, by
utilizing a GFP-tagged strain of P. ananatis, the colonization of onion thrips feeding injury by P.
ananatis was observed using fluorescence microscopy. These studies suggests that onion thrips
play a significant role in the development of bacterial leaf blight and center rot in onions. Results
from the pesticide trial indicate that thrips feeding damage is positively correlated with bacterial
leaf blight incidence. Therefore, in order to improve bacterial disease suppression in onions,
management efforts should focus on reducing onion thrips populations.

This thesis is dedicated to my mother and father
Thank you for your unconditional support

iii

ACKNOWLEDGMENTS

First and foremost, I would like to thank my advisor Zsofia Szendrei for her guidance
during my program. Her support and direction has been essential to the completion of this thesis
and my development as a scientist. I would also like to thank the other members of my
committee, Mary Hausbeck and Edward Walker. Mary has been a vital source of counsel,
perspective and technical expertise. Ned was equally invaluable and provided me extra
opportunities to develop new molecular skills which I will continue to use throughout my career.
Prissana Wiriyajitsomboon and Elizabeth Brisco-McCann were essential to the completion of
this work. I thank them both for their time and hard work sampling fields and leading all of the
plant pathology work included in this thesis. Schicheng Chen was critical to the development of
the GFP-tagged strain of Pantoea ananatis. I thank him for contributing his time and sharing his
expertise with me. I would like to thank undergraduates C. Mccalmon, L. Komondy, G. Nagle
and A. Thiemkey for their many hours spent crawling around the muck counting thrips and their
hard work in the lab processing samples. A special thank you to David VanderZee for his
contributions to all aspects of this thesis as well as helping me with all of the molecular work. I
would like to thank all the cooperating onion growers who allowed me access to their farms for
onion sampling and conducting the pesticide trial. I thank Dr. William J. Hickey (University of
Wisconsin-Madison) for generously providing pSCH476. Thank you to the Department of
Entomology for their support during my degree program. I thank my funding sources for this
research, MSU Project GREEEN (award no. GR15-057 to ZS and MKH), NIH (award no. R37
AI21884 to EDW) and the Michigan Onion Committee. Last but not least, I would like to thank
my family and friends for their support.

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TABLE OF CONTENTS

LIST OF TABLES ........................................................................................................................ vii
LIST OF FIGURES .................................................................................................................... viii
CHAPTER 1: Biology and management of onion thrips and bacterial diseases in onion
production systems...........................................................................................................................1
Introduction ..........................................................................................................................1
Onion pests...........................................................................................................................2
Onion thrips .........................................................................................................................4
Biology.....................................................................................................................4
Life history ...............................................................................................................5
Reproduction ............................................................................................................6
Management .............................................................................................................7
Monitoring ...............................................................................................................9
Action thresholds .....................................................................................................9
Bacterial leaf blight of onion caused by Pantoea agglomerans ........................................10
Bacterial center rot of onion caused by Pantoea ananatis ................................................12
Onion thrips as disease vectors ..........................................................................................14
Thesis objectives ................................................................................................................16
CHAPTER 2: Assessing the relationship between onion thrips (Thrips tabaci) and bacterial leaf
blight in Michigan onion fields ......................................................................................................18
Introduction ........................................................................................................................18
Materials and Methods .......................................................................................................20
Thrips and leaf blight incidence .............................................................................20
Insecticide and bactericide trial ............................................................................22
Results ................................................................................................................................24
Onion thrips and leaf blight incidence ...................................................................24
Pesticide trial ..........................................................................................................26
Discussion ..........................................................................................................................28
CHAPTER 3: Onion thrips feeding promotes infection by Pantoea ananatis in onion ................32
Introduction ........................................................................................................................32
Materials and Methods .......................................................................................................34
Onion thrips colony................................................................................................34
Experiment 1 ..........................................................................................................34
Experiment 2 ..........................................................................................................37
Experiment 3 ..........................................................................................................38
Results ................................................................................................................................40
Experiment 1 ..........................................................................................................40
Experiment 2 ..........................................................................................................43
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Experiment 3 ..........................................................................................................45
Discussion ..........................................................................................................................48
CHAPTER 4: Conclusions and future directions ..........................................................................50
APPENDIX ....................................................................................................................................55
LITERATURE CITED ..................................................................................................................57

vi

LIST OF TABLES

Table 2.1. Seasonal average number of thrips per leaf and bacterial leaf blight incidence in 2015
and 2016 when treated with a bactericide and/or an insecticide program………………………28
Table 3.1. The effects of onion thrips (Thrips tabaci), bacterial leaf blight (Pantoea ananatis)
and mechanical damage (five pricks using a pin) on onions. When thrips and bacteria were
present together, bacteria were either added to plants 7 days after thrips or simultaneously.
Thrips (3-4 adults/plant) were added to plants from a laboratory colony. Bacterial (1 × 108 CFU)
solution was sprayed onto plants using a hand spray bottle. Experiments were conducted for 14
days in a growth chamber at Michigan State University, East Lansing, MI. N = 15. Numbers (16) in the second row correspond to treatment numbers for experiment 1 used in the text……..42
Table 3.2. Effects of varying numbers of onion thrips (Thrips tabaci) per plant in combination
with bacterial leaf blight (Pantoea ananatis) on onions. Thrips (0, 4, 12 or 20/plant) were added
to plants from a laboratory colony. Plants were inoculated with a liquid culture of P. ananatis (1
× 108 CFU), sprayed onto plants. Experiments were conducted for 15 days in a growth chamber
at Michigan State University, East Lansing, MI. N = 11. Numbers (1-4) in the second through
fifth rows correspond to treatment numbers for experiment 2 used in the text………………...44
Table S.1. Voucher specimens deposited at the Albert J. Cook Arthropod Research Collection
(Michigan State University). …………………………………………………………………..56

vii

LIST OF FIGURES

Figure 2.1. Mean (± SEM) proportion of thrips that screened positive for P. agglomerans using
qPCR and mean (± SEM) proportion of plants with bacterial leaf blight symptoms at each
sampling period during the 2015 (A) and 2016 (B) growing seasons. Each point represents the
average across eight fields. Uppercase letters represent mean differences in plants with bacterial
leaf blight symptoms and lowercase letters represent mean differences in thrips positive for P.
agglomerans. Points with the same letter are not statistically significant from each other……25
Figure 2.2. The proportion of plants with bacterial leaf blight symptoms correlated with the
mean (± SEM) thrips per plant in 2015 (A) and 2016 (B). The proportion of plants with bacterial
leaf blight symptoms correlated with the proportion of thrips that screened positive for P.
agglomerans in 2015 (C) and 2016 (D). Each point represents one field for each date……….26
Figure 3.1. Correlation between the number of onion thrips (Thrips tabaci) per plant and the
percent leaf area with leaf blight symptoms (A). Correlation between the percent leaf area with
thrips feeding damage and the percent leaf area with leaf blight symptoms (B). Data are all
treatments from Experiment 2 which received thrips (treatments 2, 3 and 4; 4 – 20 thrips/plant).
Plants were inoculated with a liquid culture of P. ananatis which was sprayed onto plants (1 ×
108 CFU). Experiments were conducted in a growth chamber at Michigan State University, East
Lansing, MI. N = 11……………………………………………………………………………45
Figure 3.2. Representative samples of undamaged onion leaf tissue (A), onion tissue damaged
by thrips (Thrips tabaci) feeding (B), necrotic leaf tissue damaged by the GFP strain of P.
ananatis (C) and onion leaf tissue damaged by both thrips feeding injury and the GFP strain of P.
ananatis (D). White arrows (a) indicate where thrips mandible was inserted into the leaf tissue.
Yellow pigmentation is the result of chlorophyll removal by thrips feeding (B, D). Fluorescent P.
ananatis colonizing intercellular spaces of leaf cells (C) and regions where cells were damaged
by thrips feeding (D), indicated by the white circles (b and c, respectively). Plants were
inoculated with a liquid culture of the GFP strain of P. ananatis which was sprayed onto plants
(1 × 108 CFU). Experiments were conducted in a growth chamber at Michigan State University,
East Lansing, MI. Photos were taken at 10x magnification and each square of the reticle = 50
µm2. The scale bar = 100 µm.………………………………………………………………….47

viii

CHAPTER 1:
Biology and management of onion thrips and bacterial diseases in onion production
systems

Introduction
The importance of onion (Allium cepa L.), commonly known for causing tears in the
kitchen, as a food source for humans has been evident throughout the ages. First cultivated in
ancient Egypt (ca. 3000 BCE), onions symbolized eternity and were often found buried in the
tombs of Pharaohs (Fritsch and Friesen 2002). During modern times, onions are grown across the
world and are a staple vegetable used as an ingredient in dishes across a wide diversity of
cultures (Griffiths et al. 2002). Dry bulb onions became a staple food partly due to their longterm storage characteristics and durability during the shipping process (Griffiths et al. 2002).
Onions are grown internationally by approximately 170 countries, the leaders being
China (~33% global production), India (~13%), United States (~5%), Turkey (~3%) and
Pakistan (~3%) (FAOSTAT 2014). In the United States this industry is valued at $1 billion
dollars per year (USDA/NASS 2015). The leading producers of onions in the United States are
Washington, Idaho - Eastern Oregon and California, which combined represent approximately
50% of United States onion production (National Onion Association 2017). Michigan’s onion
industry is valued at approximately $10 million per year accounting for 1% of national onion
production (USDA/NASS 2015). In terms of acreage, Michigan currently ranks 11th in terms of
acres harvested; however, this number has been decreasing annually, falling from 5,067 acres in
1997 to 2,400 acres in 2016 (USDA/NASS 1997, 2016).

1

Onions vary in color, flavor profile, size and seasonal availability. Yellow onions make
up over 87% of the US crop while the other 13% is comprised of red and white onions (National
Onion Association 2017). Onion varieties can also be regionally specific; for example, Vidalia
onions are grown on only 6,000 hectares in Georgia but represent 40% of the spring onion
production value of the United States (Boyhan and Torrance 2002). Most of Michigan’s 45
million kilograms of commercial onions produced per year are for dry bulbs stored through the
winter, and are direct seeded.

Onion pests
Onions are attacked by a variety of viruses, bacterial and fungal diseases and insect pests.
The most common virus that affects onion is Iris Yellow Spot Virus (Bunyaviridae, Tospovirus
spp.; IYSV). IYSV was originally identified in Brazil (Pozzer et al. 1994) but has spread to other
major onion producing areas of the world (Hall et al. 1993, Gent et al. 2006). Symptoms of
IYSV appear as lesions along the edges of leaves (Gent et al. 2006) which reduce bulb size (Gent
et al. 2004) and cause yield loss (Pozzer et al. 1999). The only confirmed vector of this pathogen
is the onion thrips (Thrips tabaci Lindeman, Thysanoptera: Thripidae), an economically
important indirect pest of onion (Kritzman et al. 2001).
Many fungal pathogens cause onion diseases which affect different portions of the plant.
Pink root, caused by Setophoma terrestris (H.N. Hansen) Gruyter, Aveskamp and Verkley, is
characterized by the pink color of infected roots, and causes the shrinking and withering of onion
roots (Sumner 1997). This leads to earlier development of onion bulbs which results in a
reduction of bulb size (Sumner 1997). Purple blotch is a foliar disease of onion caused by the
fungal pathogen Alternaria porri (Ellis) which enters the plant through openings in the leaves

2

(Miller 1983). Purple blotch infection can lead to premature defoliation and can compromise
bulb quality. The feeding damage of onion thrips has been linked with purple blotch incidence,
providing alternative entry points for A. porri (McKenzie et al. 1993). Downy mildew, caused by
the fungal pathogen Peronospora destructor, causes premature death of onion leaves and a
reduction in bulb size, it can also indirectly cause storage rot (Scholten et al. 2007).
Bacterial diseases of onion cause variable levels of yield losses, and can be especially
severe when weather conditions favor the growth of pathogens. Xanthomonas leaf blight is a
common foliar disease of onion, caused by Xanthomonas axonopodis pv. allii. Xanthomonas leaf
blight favors conditions of high humidity and high temperatures (> 26 ºC) (Alvarez et al. 1978).
This disease reduces photosynthetic areas leading to a reduction in bulb size and severe yield
losses (> 20%) (Sanders et al. 2003). Two species belonging to the genus of Pantoea are causal
agents of bacterial leaf blight (Pantoea agglomerans) and bacterial center rot (Pantoea
ananatis), which; can cause foliar damage as well as bulb rot during long-term storage (Gitaitis
and Gay 1997, Edens et al. 2006). The symptoms of center rot are white necrotic streaks that run
along the length of the leaf which can progress into the bulb and predispose it to rotting caused
by secondary microbes (Gitaitis et al. 2003, Edens et al. 2006). Bacterial diseases are spread
around fields during wind and rain events (Wiriyajitsomboon et al. 2014), and often enter plant
tissue through wounds caused by extreme weather, machinery and field workers (Gitaitis et al.
1978). Onion thrips can also transmit the bacterial pathogens, P. ananatis and P. agglomerans, to
onion through feces (Dutta et al. 2014).
Onions have a variety of insect pests, both direct and indirect, that cause damage to
different areas of the plant. In the United States, the two major insect pests of onion are the onion
maggot (Delia antiqua Meigen, Diptera: Anthomyiidae), which causes damage to the bulb and

3

seedlings, in the larval stage (Whitfield et al. 1985), and onion thrips, which cause damage to
leaf tissue as both larvae and adults (Gill et al. 2015). Delia antiqua is a direct pest of alliums
since it attacks the marketable portion of the crop, the bulb (Ellis and Eckenrode 1979). It has
multiple generations in Michigan but first generation are of most concern because they cause
severe damage early in the season (Hoepting et al. 2004). Eggs, which are laid in the soil
surrounding the bulbs, hatch and larvae feed continuously on developing seedlings and bulbs (23 weeks) until they pupate into adults (Loosjes 1976). Feeding damage from D. antiqua varies
depending on the stage of the onion. In the seedling stage feeding damage results in the death of
the seedling which can reduce stands in the early season. If feeding occurs in the mid or late
season the damage results in a rotting bulb making onions unmarketable (Hausmann and Miller
1989). Since seed treatments are currently an effective way to manage D. antiqua, in Michigan,
onion thrips are the most important insect pest of onions requiring frequent insecticide
applications to maintain their populations below an economic threshold (Fournier et al. 1995).
Onion thrips has also been linked to the transmission of plant pathogens as both a direct and an
indirect vector which will be the focus of this thesis.

Onion thrips
Biology. Since its first description in 1888 (Lindeman 1889), onion thrips have been the subject
of considerable research and extension. Onion thrips possess unique asymmetrical mouthparts
made up of a single mandibular stylet and paired maxillary stylets. They use this single mandible
to pierce through the cell membrane upon which they retract the mandible and insert their
maxillary stylets (Chisholm and Lewis 1984). Pre-digestion substances are released into the plant
which help break down leaf tissue (Boateng et al. 2014). Onion thrips use a tube formed by the

4

maxillary stylets to draw up and consume mesophyll cells by cibarial pumping (Chisholm and
Lewis 1984). This unique feeding style is often referred to as “punch and suck” (Lewis 1991).
Thrips damage can be characterized by silvery leaf spots that turn into white blotches along the
leaves, a result of the loss of chlorophyll from feeding (Bailey 1938). Thrips feeding stresses
onion plants through the destruction of the leaf tissue ultimately reducing the photosynthetic
ability of the plant (Molenaar 1984, Parrella and Lewis 1997). In severe cases damage by onion
thrips has caused yield reductions of up to 60% in bulb weight (Waiganjo et al. 2008, Kendall
and Capinera 1987, Fournier et al. 1995, Rueda et al. 2007, Diaz-Montano et al. 2010). The
feeding damage caused by onion thrips can also create an entry point for plant pathogens (Orloff
et al. 2008).

Life history. Onion thrips are holometabolous insects with a lifecycle consisting of an egg, first
and second instar stages, prepupa, pupa, and adult (Ghabn 1948, Nakahara 1991, Diaz-Montano
et al. 2011). They feed on plant tissue as first and second larval instars as well as adults. Eggs are
laid singly and inserted directly into the leaf tissue. They are microscopic, averaging 0.23mm
long and 0.08mm wide (Patel et al. 2013) and incubate for a period of 4 to 5 days inside the leaf
before hatching (Fekrat et al. 2009). Larvae are semitransparent and dull white as first instars
later becoming more yellow as second instars (Patel et al. 2013, Gill et al. 2015). The average
length of first and second instars is 0.35-0.38mm and 0.7-0.9mm, respectively (Gill et al. 2015),
and they spend 2 to 3 days in first instar stage and 3 to 4 days in the second instar stage (Pourain
et al. 2009).
During the prepupal and pupal stage onion thrips do not feed. Pupation normally occurs
at the base of the onion’s apical meristem or within the soil (Rueda and Shelton 1995). The entire

5

pupal period lasts between 3 to 10 days (Patel et al. 2013), with temperature causing variation in
pupation time (Diaz-Montano et al. 2011). Adults are generally 1.0-1.3mm in length (Orloff et
al. 2008), and have a longevity varying between 28-30 days on onion (Patel et al. 2013). The
adult’s body color varies with temperature from yellow to brown (Murai and Toda 2002), and
adults possess fringed forewings and hindwings making them more mobile than larval and pupal
stages (Gill et al. 2015). Adult onion thrips are able to disperse various distances ranging from
trivial dispersal (short-range movement between plant hosts) to long-distance dispersal (longrange flight away from immediate plant hosts). Onion thrips flight tends to occur when daylight
is present, temperatures are above 17 ºC and wind speeds are low (Smith et al. 2016). Longdistance movement often occurs in the late season when insecticide applications are reduced
which can lead to the spread of plant pathogens or insecticide-resistant alleles (Smith et al.
2015).

Reproduction. Onion thrips have the ability to reproduce asexually through parthenogenesis.
Most commonly onion thrips reproduce through a mode of parthenogenesis in which females are
produced from unfertilized eggs, called thelytoky. An alternative mode of parthenogenic
reproduction, arrhenotoky, in which males are produced from unfertilized eggs, is much less
common (Gill et al. 2015). Consequently male onion thrips are much less common in most areas
of the world (Lewis 1973, Kendall and Capinera 1990). Asexual reproduction leads to multiple
problems: (1) thelytokous individuals do not need to find a mate and therefore reproduction is
not mate-limited and, (2) if the adult female is resistant to an insecticide, her offspring will be
resistant as well (Diaz-Montano et al. 2011). This becomes problematic to onion growers
because it can result in large resistant populations and major/significant economic yield losses.

6

Onion thrips development and reproduction is dependent on environmental conditions,
such as temperature. Six to eight generations per growing season have been estimated for New
York (Hoffmann et al. 1996), and similar generation numbers were observed in Kentucky and
North Carolina (Bessin 2004 and Carter and Sorenson 2013). Females begin laying eggs as soon
as temperature increases during the spring (Sites and Chambers 1990), and a 1 week
preoviposition period is followed by up to 3 weeks of egg laying (Gill et al. 2015). Higher
temperatures lead to shorter generation times (17 days at 30 ºC) and increased oviposition rates
(8 eggs per day at 25 ºC); however, the survival rate of eggs decreases at high temperatures
(10.5% at 30 ºC) (Murai 2000) . The optimal temperature for onion thrips in terms of fecundity is
approximately 23 ºC, which is a result of increased longevity of females that lay more eggs and
have low mortality (Murai 2000). Gravid females lay eggs in the same areas that they feed and
continually move toward younger leaf tissue (Theunissen and Legutowska 1991). Hot and dry
weather leads to shorter generation time and rapid population increases (Rueda et al. 2007),
which in addition to the higher rates of metabolism might also be due to factors such as a
reduction in mortality from rain and pathogens (Gill et al. 2015).

Management. Many methods of control exist which reduce the damage caused by onion thrips.
Onion cultivars which are less attractive to onion thrips include those with a yellow-green color
and glossy to semi-glossy leaf surfaces; however, no highly resistant onion cultivars exist (DiazMontano et al. 2010, Diaz-Montano et al. 2012a, Diaz-Montano et al. 2012b, Damon et al.
2014). Field location is also important when trying to manage onion thrips populations. Crops
such as alfalfa and small grains often host overwintering populations of onion thrips and should
be taken into account when choosing a field location (Carter and Sorenson 2013). Intercropping

7

of onions with other crops such as carrots (Daucus carota L.) and tomato (Solanum lycopersicum
L.) has been shown to reduce onion thrips infestations by over 50% (Uvah and Coaker 1984,
Afifi and Haydar 1990). While there are known predators of onion thrips, biological control was
determined not to be an effective means of control (Parrella and Lewis 1997, Diaz-Montano et
al. 2011, Gill et al. 2016). Predators of onion thrips; such as, Aelothrips spp., green lacewing
(Chrysoperla spp.) larvae, coccinellids (e.g. Coleomegilla maculata), big-eyed bug (Geocrois
spp.) and minute pirate bug (Orius spp.), usually do not reach high abundances until late in the
growing season, after the majority of feeding has occurred (Fok et al. 2014).
Traditionally chemical control has been used as the primary means of controlling onion
thrips and dates back to first reports of naphthalene use in 1933 (Maughan 1933 and DiazMontano et al. 2011). However, because of their short life cycle, high fecundity and asexual
mode of reproduction, onion thrips are likely to develop resistance to insecticides. Resistance to
pyrethroid and organophosphate insecticides by onion thrips populations has been reported in
onion growing areas around the world (Martin et al. 2003, Shelton et al. 2003, MacIntyre Allen
et al. 2005, Shelton et al. 2006, Herron et al. 2008, Morishita 2008). This underscores the need to
manage onion thrips insecticide resistance to maintain efficacy of currently available products.
Chemical management strategies that limit the frequency of insecticide applications, rotate
insecticides classes and maintain thorough coverage to prolong the effectiveness of insecticides,
help mitigate resistance (Gill and Garg 2014). Applying a high spray volume of insecticides to
onion improves onion thrips control and concurrently adding a penetrating surfactant critically
improves the efficacy of insecticides used to control onion thrips (Nault et al. 2013).

8

Monitoring. It is recommended that onion growers monitor their fields regularly to determine the
density of Onion thrips and time insecticide applications accordingly. Adults and larvae can be
observed and counted by opening the neck of the plant (Shelton et al. 1987). The younger leaves
are a preferred feeding site for onion thrips, therefore they should be the focus area of
examination (Rueda and Shelton 1995). It is important that sampling is started early in the
season, at the 4-5-leaf stage, and plants are checked weekly throughout the growing season,
especially if weather conditions are ideal for Onion thrips development (Rueda and Shelton
1995). Other, less commonly used methods of monitoring for onion thrips include plant
dissection and plant beating (Mautino et al. 2012). Plant dissection consists of collecting plants
from the field and dissecting and examining them leaf by leaf to detect onion thrips. Plant
beating consists of beating plants into a plastic tray and counting the onion thrips in the tray.
While plant dissection and plant beating are less accurate methods of thrips sampling compared
to visual sampling they still can be used in practice to adjust the number of thrips per plant
recorded (Mautino et al. 2012).

Action thresholds. Under favorable conditions, onion thrips populations can rapidly increase due
to their short developmental time and high fecundity (Diaz-Montano 2011) making them difficult
to control. Economic action thresholds are necessary to indicate when control measures need to
be implemented (Fournier et al. 1995). The determination of economic action thresholds depends
on factors such as weather conditions (Pedigo et al. 1986) and insecticide efficacy (Nault and
Shelton 2010), therefore no all-encompassing economic threshold exists for onion thrips on
onion (Fournier et al. 1995).

9

In most cases economic injury thresholds are region specific, such as 30 thrips per plant
in California (Kuepper 2004), 1 thrips per leaf in Texas (Edelson et al. 1989) or 5 thrips per 50
plants in Auckland, New Zealand (Jamieson et al. 2012). Action thresholds may also vary with
season or weather conditions. For example, Canada has a 0.9 thrips per leaf threshold during
early and severe drought conditions and a 2.2 thrips per leaf threshold under no or slight water
deficit (Fournier et al. 1995). More recently, in New York, specific action thresholds were
determined for different insecticides’ efficacies. For example, spirotetramat (Movento),
methomyl (Lannate LV) and abamectin (Agri-Mek SC) effectively manage onion thrips at a
threshold level of 1 thrips per leaf. In contrast, spinetoram (Radiant SC) is effective at 3 thrips
per leaf (Nault and Shelton 2010, Nault and Shelton 2012). It is important to consider geographic
and environmental factors when determining an appropriate economic action threshold for onion
thrips. If proper monitoring is implemented and the correct thresholds are followed onion thrips
control can be optimized allowing growers to make fewer applications, save time and money,
and mitigate insecticide resistance development (Gill et al. 2015).

Bacterial leaf blight of onion caused by Pantoea agglomerans
Pantoea agglomerans (Beijerinck 1888) comb. nov. (Gavini et al. 1989),
formerly Enterobacter agglomerans (Beijerinck 1888) Ewing and Fife (1972), Erwinia
herbicola (Löhnis 1911) Dye 1964 or Erwinia milletiae (Kawakami and Yoshida 1920) Magrou
1937 is a gram-negative aerobic bacillus in the family Enterobacteriaceae which can be found in
many ecological niches, most of which are associated with plants (Rezzonico 2009). However,
strains of P. agglomerans are common symbionts in arthropods such as bees (Loncaric et al.
2009, Lozo et al. 2015), ants (Suen et al. 2010) and termites (Potrikus et al. 1977). There are also

10

a few reports of P. agglomerans causing infections in species of mammals or fishes (Dutkiewicz
et al. 2016). It was identified as the cause of a hemorrhagic disease in dolphin fish (Coryphaena
hippurus), after being isolated from their kidneys (Hansen et al. 1990) and it was also identified
as the potential cause of equine abortion (Gibson et al. 1982). Lastly, strains of P. agglomerans
have been associated with human infections which follow piercing or laceration of skin with
plant material such as a thorn and subsequent inoculation of the plant-residing bacteria
(Dutkiewicz et al. 2016).
Most commonly associated with plants, P. agglomerans is an epi- or endophytic
symbiont as well as a plant pathogen. In Eucalyptus (Myrtaceae), P. agglomerans strain 33.1
was shown to possess beneficial characteristics such as plant growth promotion (Ferreira 2008).
Certain strains of P. agglomerans have been identified as biocontrol agents for fire blight in pear
orchards (Özaktan and Bora 2004) and bacterial wilt in bean (Costa et al. 2002 Hsieh et al.
2005). However, P. agglomerans has been known to cause diseases in a variety of cropping
systems such as bacterial leaf blight in rice (Oryza sativa) in Korea (Lee et al. 2010), bacterial
seed and boll rot of cotton (Gossypium hirsutum) (Medrano and Bell 2007), bacterial spot
disease of Chinese taro (Alocasia cucullata) in Brazil (Romeiro et al. 2007), brown apical
necrosis of walnut (Juglans regia) in China (Yang et al. 2011), leaf blight and vascular wilt in
maize (Zea mays) and sorghum (Sorghum bicolor) in Mexico (Morales-Valenzuela et al. 2007)
and bacterial leaf blight and center rot of onion (Dutta et al. 2014).
As a pathogen of onion, P. agglomerans was originally described as Erwina herbicola,
and was initially identified as a pathogen affecting onions in South Africa (Hattingh and Walters
1981). The first US report of this disease came from Georgia, in 2006, and symptoms were
described as water-soaked margins of the leaves and stalk, or necrotic tissue (Edens et al. 2006).

11

In 2011, P. agglomerans was reported in onion fields in six counties across Michigan (Tho et al.
2011), and is part of a disease complex, the ‘center rot complex’, with another gram-negative
bacterium, Pantoea ananatis, which has been linked to bacterial bulb rot during storage. During
its first reported year in Michigan, bacterial leaf blight incidence progressed throughout the
season and reached an incidence of 80% or greater (Tho et al. 2011), causing concern for
growers.

Bacterial center rot of onion caused by Pantoea ananatis
Pantoea ananatis is a gram-negative, aerobic, rod-shaped and yellow-pigmented bacteria
in the family Enterobacteriaceae (Coutinho and Venter, 2009). First identified in The
Philippines, P. ananatis was first discovered as the causal agent of fruitlet rot in pineapple
(Serrano 1928). Originally it formed part of the Erwinia herbicola–Enterobacter agglomerans
complex, but in 1989 it was assigned to the genus Pantoea (Gavini et al. 1989).
Similar to P. agglomerans, P. ananatis has a cosmopolitan distribution and has been
found in a variety of environments across the globe (Weller-Stuart et al. 2017). As a saprophyte,
P. ananatis has been isolated from soil, freshwater and even aviation fuel tanks (Gasser et al.
2012, Pileggi et al. 2012, Rauch et al. 2006). Pantoea ananatis has been associated with a
plethora of insects, such as fleas and ticks (Murrel et al. 2003), brown leaf hoppers (Watanabe
1996), tobacco thrips (Wells et al. 2002, Gitiatis et al. 2003, Dutta et al. 2016) and onion thrips
(Dutta et al. 2014). It was also responsible for causing bacteremic infection in a 73-year-old man
(De Beare et al. 2004).
Pantoea ananatis; however, is more commonly associated with plants and has been
isolated from a variety of cropping systems as an epiphyte, endophyte or plant pathogen. Many

12

associations of P. ananatis as an epiphyte of economically important plant species, including
wheat heads (Legard et al. 1994), Mulberry (Takahashi et al. 1995), rice (Watanabe et al. 1996),
cotton lint (Chun and Perkins 1997), poplar trees (Zeng et al. 1999), barley, buckwheat (Coplin
and Kado 2001) and maize (Paccola-Meirelles et al. 2001), have been identified. It is also a
known epiphyte of 25 asymptomatic weed species, including crabgrass, sicklepod and yellow
nutsedge (Gitaitis et al. 2002). As an endophyte, P. ananatis has been isolated from coffee
(Nunes and de Melo 2006), ginseng (Cho et al. 2007), pepper (Kang et al., 2007) and papaya
(Thomas et al., 2007). Pantoea ananatis has been identified as a plant pathogen where is can
cause diseases, such as brown rot of pineapple (Serrano 1928), graywall of tomato (Stall et all
1969), leaf blotch of sudangrass (Azad et al. 2000), leaf blight and bulb decay of onion (Gitaitis
and Gay 1997, Schwartz and Otto 2000), bacterial blight of Eucalyptus species (Coutinho et al.
2002), internal fruit rot of netted melon (Kido et al. 2008) and leaf spot of maize (Pérez-y-Terrón
et al. 2009).
As a pathogen of onion, P. ananatis was first identified in Georgia where crop losses due
to bleached leaves and rotted bulbs were reported (Gitaitis and Gay 1997). The symptoms of this
disease include white streaks and water-soaked margins along the length of the leaf that turn
necrotic as the disease progresses. The bacterium may progress further and infect the bulb,
promoting colonization by secondary microbes which results in rotting of the onion bulb (Gitaitis
and Gay 1997). Yield losses due to bacterial center rot may be as high as 100% under favorable
conditions (Gitaitis et al. 2002).
The primary source of inoculum for P. ananatis and P. agglomerans is unknown,
however there have been a few hypotheses. In Georgia, P. ananatis resides as a common
epiphyte on more than 20 weed species in onion fields (Gitaitis et al. 2002). Thus, it has been

13

suggested that weed hosts may be an inoculum source for P. ananatis. Another hypothesis is that
both P. ananatis and P. agglomerans are transmitted via seeds (Walcott et al. 2002); vertical
transfer of endophytic P. agglomerans from eucalyptus seeds to seedlings has already been
demonstrated (Ferreira et al. 2008). In the case of onions, dissemination of these bacteria usually
occurs by wind and splashing water, however, tobacco thrips (Frankliniella fusca (Hinds)) and
onion thrips are able to obtain and transmit these pathogens (Gitaitis et al. 2003 and Dutta et al.
2014).

Onion thrips as disease vectors
Onion thrips are known vectors of a variety of plant viruses and diseases; they are the
only known vectors of the tospovirus ISVY (Cortês et al. 1998, Ngata et al. 1999, Pozzer et al.
1999, Kritzman et al. 2001, Hsu et al. 2010). Their feeding injury has also been associated with
the spread of purple blotch caused by the fungal pathogen A. porri (McKenzie et al. 1993) which
uses areas of insect damage as alternative penetration sites and can cause infection on younger
leaves that are injured by onion thrips (McKenzie et al. 1993). Recently onion thrips were also
identified as a vector of P. agglomerans and P. ananatis (Gitaitis et al. 2003, Dutta et al. 2014).
Interactions between thrips and plant-pathogenic bacteria, specifically of the family
Enterobacteriaceae, are poorly understood and until recently the ability of thrips to acquire and
transmit P. ananatis, P. agglomerans and other related pathogens has not been studied. However,
recently two published studies documenting interactions between thrips species and
Enterobacteriaceae bacteria described the mechanisms of transfer, acquisition and transmission
rates of P. agglomerans and P. ananatis by onion thrips and the interactions between P. ananatis
and tobacco thrips are now understood (Dutta et al. 2014, Dutta et al. 2016).

14

Onion thrips are able to acquire both P. agglomerans and P. ananatis after just one hour
of feeding on infected onion tissue with a mean acquisition percentage of 20% for P. ananatis
and 12% for P. agglomerans. However, after 48 hours of feeding, greater than 90% of onion
thrips adults acquire these pathogens (Dutta et al. 2014). Similar results were observed for
tobacco thrips and P. ananatis (Dutta et al. 2016). After one hour of feeding on infected plants
both adults (7.1%) and larvae (4.2%) were able to acquire P. ananatis and after 48 hours >70%
of adults and larvae acquired the bacterium (Dutta et al. 2016). Results also demonstrated that
onion plants which were fed on by infected onion thrips developed bacterial leaf blight and
center rot symptoms. Isolations confirmed the identities of the bacteria as P. agglomerans and P.
ananatis (Dutta et al. 2014). In summary, onion thrips and tobacco thrips are able to vector P.
agglomerans and P. ananatis in onion under laboratory conditions (Dutta et al. 2014, Dutta et al.
2016).
To investigate the mechanism of transfer, researchers used immunolocalization to
determine distribution of P. ananatis in onion thrips and tobacco thrips (Dutta et al. 2014, Dutta
et al. 2016). When the sections of onion thrips and tobacco thrips were immunolabeled with
primary and secondary antibodies and observed under a fluorescent microscope, P. ananatis was
visible in the oesophagous, midgut and hindgut. This suggests that bacteria are only present in
the digestive tract, but not in the salivary glands or other organs. Lastly, by comparing onion
plants inoculated with onion thrips feces rinsates to plants inoculated with salivary secretions
they were able to determine a stercorarial (via feces) method of transmission (Dutta et al. 2014).
With repeated evidence of trans-stadial persistence of enterobacteraceae bacteria in thrips
it is important to determine whether thrips fitness is affected by this relationship. Dutta et al.
(2016) investigated whether or not P. ananatis affected the fecundity, survival and

15

developmental time of tobacco thrips by monitoring the time to complete a generation (adult to
adult) and adult development on surface sterilized peanut leaflets with and without epiphytic
populations of P. ananatis. There were no differences between adult survival and developmental
time between treatments which contained P. ananatis and those that did not. Results of these
experiments indicated that the presence of epiphytic populations of P. ananatis had no effect on
thrips development (Dutta et al. 2016).
My thesis research is the first investigation of the relationship between onion thrips and
pathogenic Pantoea agglomerans in the field. Onion thrips are able to acquire and transmit P.
ananatis and P. agglomerans on onions in the laboratory (Dutta et al. 2014) but, evidence of this
relationship in the field was lacking. In addition, the effect onion thrips have on the spread of
bacterial diseases in onion fields is unknown and concern from growers has been increasing ever
since bacterial leaf blight was first reported in Michigan in 2011 (Tho et al. 2011). If a vector
relationship exists in the field between onion thrips and P. agglomerans it may be more
important to focus management on the insect instead of the bacteria. An understanding of the
relationship between onion thrips and P. agglomerans in a commercial field setting is paramount
to achieving proper control of this issue.

Thesis objectives
First, I determined the role of onion thrips as a vector of P. agglomerans in commercial
onion fields in Michigan using real time polymerase chain reaction (qPCR) to evaluate the
presence of the disease in field-collected onion thrips. Secondly, I assessed how the temporal
dynamics of onion thrips and P. agglomerans are related in the field. I also investigated the
effectiveness of combining insecticides and bactericides for managing both onion thrips and P.

16

agglomerans in the field. Lastly, I determined the role of feeding injury by onion thrips in
influencing the pathogenicity of P. ananatis.

17

CHAPTER 2:
Assessing the relationship between onion thrips (Thrips tabaci) and bacterial leaf blight in
Michigan onion fields

Introduction
Pantoea agglomerans (syn. Erwinia herbicola; causal agent of bacterial leaf blight)
results in symptoms of leaf blight on onion, Allium cepa L. (Amaryllidaceae), including necrotic
streaking with water-soaked margins along the length of the leaf leading to wilting in severe
cases (Edens et al. 2006). Leaf blight caused by P. agglomerans produces symptoms similar to
those by Pantoea ananatis (causal agent of center rot); thus P. agglomerans is considered part of
the onion center rot ‘complex’ (Edens et al. 2006). Bacterial leaf blight is a cosmopolitan pest,
first reported in onions in South Africa in 1981 (Hattingh and Walters 1981). In the United
States, this disease was first found in the field in Georgia in 2006 (Edens et al. 2006) and, in
Michigan, it was confirmed on onions in six counties in 2013 (Tho et al. 2015).
Onion thrips, Thrips tabaci Lindeman 1889 (Thysanoptera: Thripidae), is an important
pest of onion and many other crops worldwide (Lewis 1997). The larvae and adults feed on leaf
tissue, removing chlorophyll, thus impairing photosynthesis (Molenaar 1984, Parrella and Lewis
1997), ultimately reducing bulb size and causing yield losses (Fournier et al. 1995). In Michigan,
as in many other places in the world, onion thrips are the main insect pest of onions and have
been directly and indirectly associated with the transmission of several onion pathogens,
emphasizing the need for effective management (Diaz-Montano et al. 2011). Insecticides are
commonly used for onion thrips management (Diaz-Montano et al. 2011) but onion thrips have
become resistant to many different chemistries (Lewis 1997). Growers are advised to rotate

18

insecticides classes within a growing season and apply them based on established thresholds
(Byrne and Szendrei 2013).
Onion thrips are known to play an important role in the transmission of a plant
pathogenic virus, Iris yellow spot virus (Hsu et al. 2010, Kritzman et al. 2001, Pozzer et al.
1999), and a pathogenic fungus that causes purple blotch (McKenzie et al. 1993). Laboratory
experiments confirmed that onion thrips can acquire and transmit both P. ananatis and P.
agglomerans, and that the mode of transmission of P. ananatis was stercorarian (Dutta et al.
2014). Tobacco thrips (Frankliniella fusca (Hinds)) can also transmit P. ananatis, and while the
pathogen can persist through multiple life stages, it has no effect on tobacco thrips fitness (Dutta
et al. 2016). Based on these laboratory findings, onion thrips act as a vector for P. agglomerans,
but field research is needed to determine whether there is a link between onion thrips and leaf
blight incidence in onions.
In the United States the onion industry is valued at $1 billion dollars per year
(USDA/NASS, 2015). Michigan’s onion industry is valued at approximately $10 million per
year, grown on about 1,000 hectares, producing just over 45 million kilograms per year
(USDA/NASS 2015). Most of Michigan’s commercial onions are produced for dry bulbs stored
through the winter, and are direct seeded. In Michigan, bacterial leaf blight incidence can reach
up to 80% or more in some commercial onion fields (Tho et al. 2015). The long-term effects of
P. agglomerans on the onion industry are unknown, but with its association to other center rot
causing bacteria, there is potential for it to cause economic losses (Dutta et al. 2014). Yield
losses caused by onion thrips feeding are variable but have been documented as high as 60%
(Waiganjo et al. 2008).

19

The objectives of this study were to determine whether there is a relationship between
onion thrips and bacterial leaf blight incidence in the field and to test the efficacy of insecticides
and bactericides in limiting onion thrips and bacterial leaf blight.

Methods and Materials
Thrips and leaf blight incidence. Onion plants from two fields from each of four Michigan
commercial onion farms located in Newaygo, Allegan, Ingham and Clinton counties were
sampled in 2015 and 2016 (N=8 per year). All of the selected fields had muck soils and field
sizes ranged from 3.0 - 4.5 hectares. For each of the eight fields in both seasons, ten sampling
plots (7.62 m2 each) were established per field.
Onion foliage was sampled for thrips and bacterial leaf blight symptoms weekly between
15 June and 10 August 2015 (nine sampling dates) and 6 June and 8 August 2016 (ten sampling
dates). Plants were between 2 and 4 leaf stage when sampling began; five onion plants were
randomly collected from each of the ten plots for a total of 400 onions collected per week across
all fields. Individual plants were removed from the soil, placed singly in plastic bags in order to
retain all thrips and placed into coolers for transport to the laboratory.
Each sample was assessed for thrips by counting the total number of adults and juveniles.
Between 1 and 15 thrips per plant were collected to screen for P. agglomerans. Insects selected
for DNA extraction were frozen in 90% ethanol.
Whole thrips DNA extractions were performed using the DNeasy blood and tissue kit
(Qiagen, Hildon, Germany). The protocol was modified by using 25% of all extraction buffers to
optimize the concentration of DNA (25.0 ± 3.0 ng / µl) in samples containing 1 to 15 thrips that
were eluted with 50 µl buffer. Extracted DNA was screened for the presence of P. agglomerans

20

using quantitative PCR (qPCR) with the species specific pagRrt2 primer set and pagR2 probe
(Braun-Kiewnick et al. 2012). qPCR reactions were performed in final reaction volumes of 10 µl
containing 2 x 5 µl TaqMan master mix (Life Technologies, Grand Island NY), 0.4 µl ddH20,
900 nM pagRrt2 forward and reverse primers, 200 nM pagR2 TaqMan probe and 2 µl extracted
DNA. PCR reactions were performed on a StepOnePlus™ Real-Time PCR System (Applied
Biosystems®, Foster City, CA). Cycling conditions were 2 min at 50 ºC, 10 min at 95 ºC,
followed by 45 cycles of 15 s at 95 ºC and 2 min at 60 ºC. All PCR reactions were conducted as
duplicates of each sample. Any sample with an average cycle threshold (CT) value < 40 was
considered positive (Braun-Kiewnick et al. 2012).
Plants were visually assessed for the presence or absence of bacterial leaf blight
symptoms. A single symptomatic plant was randomly selected from each plot for bacterial
isolation and identification. Plants were rinsed with running tap water and the foliage was
allowed to dry for approximately 1 hr. Leaf tissue from the margin of the water-soaked tissue
was excised (1.0 x 1.0 cm2) using a sterile scalpel blade, dipped briefly in 95% ethanol, rinsed
with sterile distilled water twice, dried for 3 min, and placed on a clean microscope slide. The
disinfected leaf piece was cut into half using a sterile scalpel blade. A 50 µl drop of sterile
distilled water was pipetted onto the cut area where bacterial cells were released from the tissue
into the water. The suspension was streaked onto nutrient broth yeast extract (NBY) agar using a
sterile bacterial loop, and incubated at 30⁰C for 2 to 7 days. Each bacterial colony that had
different morphological characteristics (form and color) was selected and transferred to fresh
NBY agar twice, resulting in pure cultures for identification. Representative colonies were
identified to species using the BIOLOG GN Microtiter Plates (BIOLOG Inc., Hayward, CA,
USA) according to the manufacturer’s instructions.

21

In 2015, selected samples were further confirmed via sequencing using the following
protocol: DNA extraction was performed according to DNA extraction manual using the Wizard
Genomic DNA Purification kit (Promega Corp., Leiden, The Netherlands). DNA quantity was
measured using Nanodrop 1000 spectrophotometer (Thermo Scientific, Wilmington, DE). 16S
rDNA was amplified with 16S primer set. A total volume of 25 μl of PCR mixture contained 1x
PCR buffer, 0.2 μM dNTPs, 0.2 μM of each primer, and 0.625 U Taq polymerase (Promega,
Madison, WI). The PCR reactions were conducted in a Mastercycler Pro thermocycler
(eppendorf, Hauppauge, NY) with an initial denaturation at 94℃ for 5 min, followed by 3 cycles
of denaturation at 94℃ for 45 sec, annealing at 52℃ for 2 min, and extension at 72℃ for 1 min,
and another 30 cycles of denaturation at 94℃ for 20 sec, annealing at 52℃ for 1 min, and
extension at 72℃ for 1 min. The final extension step was performed at 72℃ for 7 min. The
PCR products were visualized on 1.5% agarose gel, dissolved in 1x Tris-borate-EDTA (TBE)
buffer, and sequenced with a single primer by submitting to Macrogen Corp. (Macrogen USA,
Rockville, MD). The nucleotide sequences were compared to the nucleotide collection in NCBI
using a BLASTn search (http://blast.ncbi.nlm.nih.goc.Blast) analysis.
Pearson product-moment correlation coefficients were computed to assess the
relationships between mean thrips per plant, proportion of thrips positive for P. agglomerans and
proportion of plants with bacterial leaf blight symptoms. Differences among dates for each
dependent variable were compared with an analysis of variance (ANOVA). Differences between
dates were separated using a Tukeys HSD (R, version 3.2.2).

Insecticide and bactericide trial. To assess the effectiveness of different bactericides applied
alone or in combination with an insecticide, a field trial was conducted during the 2015 and 2016

22

growing seasons. ‘Sedona’ onions were direct seeded on 29 April 2015 and 16 April 2016 in a
commercial field in Stockbridge, MI. The trial was arranged in a randomized complete block
design with 13 treatments replicated across four blocks. Each plot was three rows wide and 6 m
long with a 60 cm buffer zone in between each plot. Three bactericides were tested: two copper
hydroxide products (Kocide 3000®, DuPont™, Wilmington, DE and Nu-Cop® 50, Albough Inc.,
Ankeny, IA), and an antibiotic (Kasugamycin®, Hokko Chemical Industry Co. Ltd, Tokyo,
Japan) (Table 2.1). A grower standard insecticide program was tested alone or in combination
with the bactericides and a non-ionic surfactant (Dyne-Amic®, Helena Chemical Company,
Collierville, TN) (Table 2.1). In 2015, the insecticide program consisted of two applications of
spinosad (Radiant® SC, Dow™ AgroSciences, Indianapolis, IN) followed by two applications of
abamectin (Agri-Mek® SC, Syngenta®, Basel, Switzerland) at weekly intervals. In 2016, the
insecticide program consisted of weekly sprays consisting of the following: two applications of a
spirotetramat (Movento®, Bayer CropScience LP, RTP, NC), followed by two applications of
spinosad, followed by two applications of abamectin followed by one application of methomyl
(Lannate® LV, DuPontTM, Willmington, DE). A penetrating surfactant (Dyne-Amic, Bayer
CropScience LP) was added at 0.05% v/v in some treatments (Table 2.1). These treatments were
compared to an untreated control. The bactericide program was initiated on 28 May 2015 and 22
June 2016 when plants had 3 to 5 leaves. The insecticide program was based on thrips/leaf
thresholds (Nault and Shelton 2010) and was initiated on 22 July 2015 and 29 June 2016. All
combination treatments were tank mixed before application. All products were applied using a
CO2 backpack sprayer and a broadcast boom equipped with three XR8003 flat-fan nozzles
(TeeJet® Technologies, Wheaton, IL) with the outer nozzles angled towards the center of the
plot, calibrated at 50 psi and delivering 467.5 L ha-1.

23

Prior to initiating insecticide applications, plants were scouted weekly and the number of
thrips per plant was used to calculate the thrips per leaf threshold. After the insecticide program
began, thrips were counted on ten randomly selected plants in each plot weekly. The total
number of leaves per plant was also counted on twenty randomly selected plants in the field. The
average number of thrips per leaf for each treatment was calculated.
Leaf necrosis was visually evaluated according to the Horsfall-Barratt rating system
where 1 indicates no disease symptoms and 12 indicates up to 100% foliar necrosis (Horsfall
Barratt 1945). Plants were evaluated one week following the final pesticide application: on 24
August in 2015 and on 17 August in 2016.
The seasonal mean number of thrips per plant was compared among the thirteen
treatments using an ANOVA where treatment was the fixed factor and block by date interaction
was the random factor. Mean necrotic leaf tissue was also compared among the thirteen
treatments using an ANOVA where treatment was the fixed factor and block was the random
factor. Tukey’s HSD tests were used to determine differences among treatment means.

Results
Onion thrips and leaf blight incidence. The number of thrips per plant, thrips (%) that tested
positive for P. agglomerans, and plants (%) with bacterial leaf blight symptoms increased during
each growing season. The mean number of thrips per plant significantly differed among
sampling dates (2015: F = 35.9, df = 8, 62, p < 0.01; 2016: F = 2.6, df = 9, 61, p < 0.01),
reaching about four thrips per plant by the end of the growing season. Thrips (%) positive for P.
agglomerans also differed among sampling dates (2015: F = 9.7, df = 8, 62, p < 0.01; 2016: F =
4.3, df = 9, 61, p < 0.05; Figure 2.1). Plants (%) with bacterial leaf blight significantly differed

24

among sampling dates (2015: F = 31.8, df = 8, 62, p < 0.01; 2016: F = 16.0, df = 9, 61, p < 0.01)
reaching 100% by the end of the growing season (Figure 2.1).

There was a significant positive correlation between plants (%) with bacterial leaf blight
symptoms and the mean number of thrips per plant in both growing seasons (2015: r2 = 0.26, p <
0.05; 2016: r2 = 0.30, p < 0.01; Figure 2. A, B). There was a significant positive correlation

25

between plants (%) with bacterial leaf blight symptoms and thrips (%) that screened positive for
P. agglomerans in both growing seasons (2015: r2 = 0.59, p < 0.01; 2016: r2 = 0.23, p < 0.05;
Figure. 2 C, D).

Pesticide trial. In both growing seasons, all treatments with an insecticide had significantly
fewer thrips than treatments without insecticide (2015: F = 57.5, df = 12, 12, p < 0.01; 2016: F =
71.4, df = 12, 24, p < 0.01; Table 2.1). In 2015, the treatments containing Kasugamycin® and
insecticide provided better control of thrips than the insecticide treatments containing the other
two bactericides (Kocide®, NuCop®). The number of thrips among treatments containing

26

bactericides mixed with insecticide compared to insecticide alone did not differ in 2015 and
2016. Adding a surfactant did not significantly impact thrips control or leaf blight severity (Table
2.1).
There was a significant treatment effect on leaf blight severity in the pesticide trial in
both growing seasons (2015: F = 6.5, df = 12, 36, p < 0.01; 2016: F = 24.8, df = 12, 36, p < 0.01;
Table 2.1). In 2015, all treatments that contained an insecticide showed 26% less plant area with
leaf blight symptoms than the treatments without insecticide, but this effect was not significant.
In 2016, all treatments with an insecticide had 35% less plant area with bacterial leaf blight
symptoms than treatments without insecticide (t > 5.2, df = 12, p < 0.01). Leaf blight severity
was similar among the eight treatments that contained an insecticide in both years (Table 2.1).

27

Table 2.1. Seasonal average number of thrips per leaf and bacterial leaf blight incidence in 2015 and 2016 when treated with a
bactericide and/or an insecticide program.
2015

2016

Seasonal avg.
Seasonal avg.
Necrotic leaf
Necrotic leaf
no. thrips/leaf
no.
thrips/leaf
y,z
tissue ± SE
tissue ± SEy,z
± SEx,z
± SEx,z
3.7 ± 0.27 a
8.0 ± 0.63 abc
8.2 ± 0.47 b
6.5 ± 0.29 a
Kocide 3000
Cu(OH)2
3.8 ± 0.27 a
8.0 ± 0.63 abc
11.5 ± 0.75 a 6.0 ± 0.00 a
Kasugamycin Kasugamycin
3.0 ± 0.21 a
8.1 ± 0.00 abc
9.9 ± 0.63 ab 6.8 ± 0.25 a
NuCop 50
Cu(OH)2
3.3 ± 0.26 a
9.2 ± 0.63 a
10.4 ± 0.82 a 6.5 ± 0.29 a
Kocide 3000
Cu(OH)2
+
1.3 ± 0.13 bcd
5.8 ± 0.25 c
4.8 ± 0.32 c
3.8 ± 0.25 b
Kasugamycin Kasugamycin
+
0.8 ± 0.08 d
7.0 ± 0.85 bc
4.2 ± 0.31 c
3.8 ± 0.25 b
NuCop 50
Cu(OH)2
+
1.4 ± 0.12 b
6.0 ± 0.41 c
4.5 ± 0.32 c
4.8 ± 0.25 b
Kocide 3000
Cu(OH)2
+
+
1.4 ± 0.13 b
5.8 ± 0.48 c
3.9 ± 0.25 c
4.5 ± 0.29 b
Kasugamycin Kasugamycin
+
+
0.9 ± 0.12 cd
6.3 ± 0.48 bc
3.2 ± 0.24 c
4.0 ± 0.00 b
NuCop 50
Cu(OH)2
+
+
1.4 ± 0.12 bc
6.3 ± 0.25 bc
3.7 ± 0.24 c
4.5 ± 0.29 b
+
1.2 ± 0.13 bcd
6.3 ± 0.48 bc
3.7 ± 0.29 c
4.3 ± 0.25 b
+
+
1.0 ± 0.10 bcd
5.8 ± 0.48 c
3.7 ± 0.27 c
4.0 ± 0.00 b
+
3.1 ± 0.21 a
8.6 ± 0.50 ab
10.2 ± 0.61 a 6.5 ± 0.29 a
v
-1
All treatments that contain copper hydroxide (Cu(OH)2) were applied at 1.68 kg ha and all treatments that contain Kasugamycin
were applied at 2.34 L ha-1.
w
Non-ionic surfactant (Dyne-Amic ®) was added at 0.05% v/v.
x
The seasonal average number of thrips per leaf was calculated by averaging the sum of thrips per leaf and dividing by the average
number of leaves per plant.
y
Necrotic leaf tissue was measured at the end of the season using the Horsfall-Barratt rating system (1 = 0-3% of necrotic leaf tissue
and 12 = 100% of leaf necrosis).
z
Different letters indicate differences among treatments, Tukey’s HSD, α < 0.0.
Product Name

Active
ingredientv

Insecticide
Program (+/-)

Surfactant
(+/-)w

28

Discussion
Results from this study indicate that onion thrips are likely contributing to the spread of
the bacterial leaf blight pathogen in onion fields. In each season, I detected bacterial leaf blight
symptoms approximately three weeks before thrips populations began to increase. Thrips
populations started increasing in late July in both growing seasons, which coincided with 90 to
100% incidence of bacterial leaf blight symptoms across all fields. Thrips that were positive for
P. agglomerans peaked at approximately 20% by the end of the sampling period, close to the
time of harvesting the bulbs. Insecticide applications are often decreased near harvest time that
can lead to an increase in thrips populations and this also coincides with exodus flights of thrips
from the field (Smith et al. 2015). The role of overwintering thrips in introducing the bacterium
to subsequent onion crops is unknown; however, the fact that 6% of the thrips samples were
positive for P. agglomerans in the first samples taken for both years indicates that this may be a
potential mechanism for the disease to survive and spread.
The field data alone cannot prove whether onion thrips are vectoring the bacterium since
the incidence of plant disease could be expected to increase during the growing season if not
managed effectively (Pullman and DeVay 1982). However, the field surveys together with data
from the pesticide trial support thrips as vectors of P. agglomerans because insecticide
treatments significantly reduced leaf blight symptoms. The incidence of P. agglomerans
throughout the field can be associated with multiple methods of thrips transfer. While
stercorarian mode of transmission has been verified (Dutta et al. 2014), it is likely that under
field conditions thrips also carry bacteria on their integument. Furthermore, it is likely that thrips
feeding provides entrance points for P. agglomerans inoculation. A similar relationship has been
documented between onion thrips and the fungal pathogen Alternaria porri, causal agent of

29

purple blotch (McKenzie et al. 1993). When thrips injury was present, A. porri was able to
penetrate through areas damaged by onion thrips. These methods did not differentiate whether
thrips were carrying bacteria in their guts or on their integument; therefore, I am not able to
distinguish the relative importance of the mode of transmission. However, P. agglomerans may
also be carried to the plants by machinery, wind and splashing water (Wiriyajitsomboon et al.
2014), then use thrips feeding injury as entry points into the plant.
Pantoea agglomerans is part of a disease complex that includes the bacterial center rot
pathogen, P. ananatis. It is important to prevent both Pantoea species from progressing into the
bulb late in the growing season because they can cause rot during long-term storage (Edens et al.
2006, Vahling-Armstrong et al. 2016). Currently, P. ananatis is not as prevalent as P.
agglomerans in Michigan’s commercial onion industry, but the similar acquisition rates and
modes of vectoring of the two bacteria indicate that onion thrips are able to spread both
pathogens in a similar way (Dutta et al. 2014, Dutta et al. 2016). Effectively managing onion
thrips populations could reduce the risk of losses from bulb rot during long-term storage.
A better understanding of P. agglomerans epidemiology is needed to further refine an
effective control strategy. For instance, Pseudomonas syringae pv. tomato (Okabe) survives on
the tomato plant only becoming symptomatic under appropriate environmental conditions
(Smitley and McCarter 1982). Understanding the population threshold that must be exceeded for
symptom development could be useful in order to time insecticide applications for thrips control
to minimize pathogen transmission and spread. Susceptibility to P. agglomerans at different
onion plant ages could also be a future avenue for research. In the case of purple blotch
(Alternaria porri), onion leaves are more susceptible to infection as they age and emerging
leaves are more susceptible in late season, closer to bulb maturity (Miller 1983).

30

The inoculum source for P. agglomerans is currently unknown but it has been previously
identified from seeds (Walcott et al. 2002), various weed species (Gitaitis et al. 2002) and a
variety of other crops such as Sundangrass (Azad et al. 2000), muskmelons (Bruton et al. 1986),
cantaloupe (Bruton et al. 1991), Eucalyptus (Coutinho et al. 2002), pineapple (Serrano 1928),
and honeydew melons (Wells et al. 2011). Onion thrips are known to feed and overwinter on a
variety of weed species (Smith et al. 2011) before colonizing volunteer onions in the field the
following spring (Larentzaki et al. 2007). If onion thrips are interacting with P. agglomerans on
other host plants before the start of the growing season, they may be introducing it into the onion
field. Screening volunteer onions and weeds early in the growing season for bacterial leaf blight
symptoms as well as thrips carrying P. agglomerans could provide important insight as to how
the pathogen is being introduced into onion fields.
This study demonstrates that increasing thrips populations are associated with increased
leaf blight in commercial onion fields in Michigan. Future research should focus on revising
action thresholds to optimize onion thrips and bacterial leaf blight management.

31

CHAPTER 3:
Onion thrips feeding promotes infection by Pantoea ananatis in onion

Introduction
Onion thrips, Thrips tabaci Lindeman (Thysanoptera: Thripidae), is a primary pest insect
of onions (Allium cepa L.), directly affecting the United States’ onion industry annually valued at
$1 billion (USDA/NASS, 2015). Feeding by onion thrips results in the destruction of epidermal
and parenchyma tissue (Chisholm and Lewis 1984, Hunter and Ullman 1989) and leads to
chlorophyll loss that reduces photosynthetic conductivity (Diaz-Montano et al. 2011, Gill et al.
2015). In severe cases, high thrips infestation rates can result in bulb weight reductions of up to
60% (Kendall and Capinera 1987, Fournier et al. 1995, Rueda et al. 2007, Waiganjo et al. 2008,
Diaz-Montano et al. 2011). In addition to directly feeding on plant tissue, onion thrips are also
known vectors of several onion diseases (Gill et al. 2015). For example, they are able to acquire
and transmit Pantoea ananatis (Serrano) Mergaert, a bacteria that causes center rot in onions
(Gitaitis and Gay 1997, Lewis 1997, Dutta et al. 2014, Dutta et al. 2016). Pantoea ananatis and
close members of the onion center-rot complex, Pantoea agglomerans (syn. Erwinia herbicola)
and Pantoea allii (Brady et al. 2011), produce leaf-symptoms with white streaks and watersoaked margins along the length of the leaf that turn necrotic as the disease progresses; they also
cause center rot during storage (Gitaitis and Gay 1997, Vahling-Armstrong et al. 2016). Yield
losses associated with center rot may be as high as 100% under favorable conditions (Gitaitis et
al. 2002).
Multiple species of thrips are able to acquire P. ananatis through feeding and transmit the
bacteria through their feces, i.e., by stercorarial means (Gitaitis et al. 2003, Dutta et al. 2014,

32

Dutta et al. 2016). Under laboratory conditions, 20% of onion thrips acquired P. ananatis after 1
hour of feeding on infected onions; 48 hours of feeding resulted in 100% acquisition (Dutta et al.
2014). Center rot symptoms were also observed on 70% onion seedlings after 15 days of feeding
by P. ananatis-infected thrips (Dutta et al. 2014). Similarly, 65% of onion seedlings inoculated
with rinsates containing P. ananatis-infected thrips feces developed center rot symptoms
indicating the P. ananatis could be transmitted from the thrips gut via feces following oral
ingestion (Dutta et al. 2014). However, this relationship has not been tested under field
conditions where more biotic and abiotic factors are at play. Recent research in Michigan
determined that thrips in commercial onion fields play a role in the transmission of Pantoea sp.,
where higher abundances of onion thrips was correlated with a higher proportion of necrotic
leaves in the onion plantings (Grode et al. 2017).
Bacterial infections, such as those caused by P. ananatis, are spread in agricultural fields
during wind and rain events (Wiriyajitsomboon et al. 2014). The bacteria frequently enter plant
tissue through mechanical wounds (Giataitis et al. 1978, Gitaitis et al. 2002). A recent field trial
demonstrated that a season-long insecticide program led to lower thrips populations as well as
reduced bacterial symptoms, regardless of whether onions were also treated with bactericides
(Grode et al. 2017). These results suggest that thrips feeding may lead to an increase in infection
by Pantoea sp. in the field. A similar relationship has been documented with onion thrips and
purple blotch, a fungal disease, where thrips feeding wounds enhanced the development of
purple blotch by providing alternative entry points for it (McKenzie et al. 1993). Similarly,
feeding wounds caused by onion thrips may provide entry points for P. ananatis, leading to
increased necrotic leaf tissue and center rot development.

33

The objectives of this study were to 1) determine the role of onion thrips feeding injury in
facilitating the development of P. ananatis, 2) investigate the relationship between thrips
abundance on plants and center rot development and, 3) quantify morphological changes in
onion tissue from onion thrips feeding and P. ananatis infection.

Materials and Methods
Onion thrips colony. Onion thrips used in these experiments were from a laboratory colony kept
in cages (50 cm by 50 cm by 50 cm; MegaView, Taichung, Taiwan) at Michigan State
University (East Lansing, MI). Insects originated from wild populations of onion thrips collected
from a commercial onion farm (Stockbridge, MI) in 2015. The colony was maintained on potted
onion plants (cv. ‘Sedona’; High Mowing Organic Seeds, Wolcott, VT) and allowed to feed ad
labitum; new plants were added every one to two weeks allowing thrips to transfer from old to
new plants. The colony was kept at room temperature (22-25 C), with ambient humidity.

Experiment 1. ‘Sedona’ onions were seeded (High Mowing Organic Seeds) in 4 cm square pots
(FarmTek, Dyersville, IA) and maintained in an environmental growth chamber (16:8 L:D, 28
ºC, 40% RH). Plants received water and fertilizer (20-20-20 N-P-K; Jack’s Professional®, J.R.
Peters, Inc., Allentown, PA) every other day. After 5 weeks, onion plants of uniform size (3-5
leaf stage) were transplanted into 12 cm square pots (The HC Companies: ITML, Middlefield,
OH). An experimental unit consisted of a plant surrounded by a ventilated cylindrical cage (11
cm tall, 2.5 cm wide) made of polyester plastic (ACCO Brands, Inc.: Apollo, Lincolnshire, IL),
embedded in soil (2-3 cm deep) with a top made of nylon netting attached with hot glue (160 µm
mesh size, MegaView). Cages prevented the movement of thrips from plant to plant but allowed

34

light and air movement into the cages. Plants in the environmental chamber were arranged in a
randomized complete block design with six treatments and 15 replications. The six treatments
were: (1) an undamaged control, (2) thrips only, (3) P. ananatis only, (4) thrips 7 days prior to P.
ananatis inoculation, (5) mechanical injury at the same time as P. ananatis and (6) thrips at the
same time as P. ananatis inoculation. Plants receiving thrips were infested with one adult
thrips/leaf (3-5 thrips/plant). All treatments receiving bacteria (treatments 3, 4, 5 and 6) were
inoculated with a virulent strain of P. ananatis (~1 × 108 CFU/ml, Dutta et al. 2014), isolated
from symptomatic onion plants grown in a commercial onion field (Stockbridge, MI).
Inoculations were done using a 250 ml spray bottle (Meijer Inc., Grand Rapids, MI) until leaves
were fully wetted, at 7 days after the initial thrips infestation. Mechanical damage was inflicted
by poking 5 holes/leaf randomly using a pin to mimic thrips feeding wounds. Plants were grown
for 7 days in an environmental growth chamber (16:8 L:D, 28 ºC, 40% RH), then returned to the
laboratory for rating, after which they were removed from soil and placed singly in ziptop bags
to retain thrips for later counting.
The number of leaves per plant and the length of each leaf was recorded prior to adding
treatments and after the 14-day treatment period. Total leaf area was calculated using leaf length
as an index of leaf area (Gamiely et al. 1991). Growth of each plant was calculated by
subtracting the total leaf area of each plant prior to thrips infestation from the total leaf area of
each plant calculated at 7 days after inoculation. The percentage of leaf area affected by P.
ananatis and thrips feeding was visually estimated for each plant (Horsfall and Barratt 1945).
Thrips damage was assessed by estimating the percentage of leaf area fed on by thrips and the
total number of adults and juveniles were counted on each plant.

35

Five symptomatic plants were randomly selected from each treatment for bacterial
isolation and identification. Leaf tissue from the water-soaked margin of the leaves was excised
(1.0 x 1.0 cm2) using a sterile scalpel blade, dipped briefly in 95% ethanol, rinsed with sterile
distilled water twice, dried for 3 min and placed on a clean microscope slide. The disinfected leaf
piece was macerated using a sterile scalpel with a # 10 blade (Bard-Parker®, Becton Dickinson
Acute Care, Franklin Lakes, NJ). A 50 µl drop of sterile distilled water was pipetted onto the cut
leaf where bacterial cells were released from the tissue into the water. The suspension was
streaked onto nutrient broth yeast extract (NBY) agar using a sterile bacterial loop, and incubated
at 28 C for 2 days. Each bacterial colony that had different morphological characteristics (form
and color) was selected and transferred once to fresh NBY agar resulting in pure cultures for
identification.
Representative colonies were identified to species by sequencing the small subunit of the
bacterial ribomosomal DNA gene. First, DNA was extracted using the DNeasy® Blood and
Tissue Kit (Qiagen®, Hilden, Germany). DNA was quantified using a Qubit® 2.0 Flourometer
and Qubit® dsDNA HS Assay Kit (ThermoFisher Scientific Inc., Waltham, MA). The 16S rDNA
gene was amplified with a 16S primer pair (forward primer, AGTTTGATCCTGGCTCAG,
reverse primer TACCTTGTTACGACTTCGTCCCA; De Baere et al. 2004). A total volume of
50 µl PCR mixture contained, 38.5 µl PCR-grade H2O, 10x PCR buffer, 0.2 µM dNTPs, 0.2 µM
of each primer, and 0.625 U Taq polymerase (Promega, Madison, WI). PCR reactions were
conducted in a Mastercycler Pro thermocycler (Eppendorf, Hauppauge, NY) with an initial
denaturation at 94 ºC for 5 min, followed by 3 cycles of denaturation at 94 ºC for 45 sec,
annealing at 52 ºC for 2 min and extension at 72 ºC for 1 min, and another 30 cycles of
denaturation at 94 ºC for 20 sec, annealing at 52 ºC for 1 min and extension at 72 ºC for 1 min.

36

The final extension step was performed at 72 ºC for 7 min. The PCR products were visualized on
1.5% agarose gel stained with 7.5 µl GelRed nucleic acid stain (Phenix Research Products,
Candler, NC). Reactions with sufficient PCR product were purified using the QIAQuick® PCR
Purification Kit (Qiagen®) and sequenced with a single primer by submitting to the Michigan
State University Genomics Core Facility (East Lansing, MI). The nucleotide sequences were
compared to the nucleotide collection in NCBI using a BLAST® nucleotide search analysis.
Mean plant growth, leaf area, number of leaves, bacterial leaf blight severity, thrips
damage and thrips per plant were compared among treatments using an analysis of variance
(ANOVA). All of the variables met the assumptions of ANOVA. Tukey’s HSD tests were used
to determine differences among treatment means. All statistical analysis was conducted using R
software (R Core Development Team, 2015).

Experiment 2. Plants in this experiment were grown and maintained as described in Experiment 1
except they were grown for 7 weeks (4-6 leaf stage). In order to measure the effects of varying
degrees of thrips feeding damage on bacterial leaf blight severity, plants were arranged in a
randomized complete block design with four treatments and 11 replications. An experimental
unit consisted of a plant surrounded by a ventilated cylindrical cage as described in Experiment
1. The four treatments included: (1) P. ananatis only, (2) 4 thrips per plant with P. ananatis, (3)
12 thrips per plant with P. ananatis and (4) 20 thrips per plant with P. ananatis.
Eight days after thrips infestation, all treatments were inoculated with a virulent strain of
P. ananatis (~1 × 108 CFU/ml) as described in Experiment 1. Onions were rated 7 days
following bacterial inoculation as described in Experiment 1.

37

Mean plant growth, leaf area, number of leaves, bacterial leaf blight severity, thrips
damage and thrips per plant were compared among treatments using an ANOVA. All of the
variables met the assumptions of an ANOVA. Tukey’s HSD tests were used to determine
differences among treatment means. Pearson product-moment correlation coefficients were
computed to assess the relationships between the number of thrips present on plants or thrips
feeding damage (%) and leaf blight severity.

Experiment 3. Plasmid pSCH476, derived from the wide-range vector PBBR1MCS-3 in Gramnegative bacteria, contained a gfp gene expression cassette and a tetracycline resistance gene. It
was successfully used for labelling Delftia sp. Cs 1-4 (Chen and Hickey 2011) and attempted in
P. ananatis.
To make the competent cells for transformation of pSCH476, the pathogenic P. ananatis
strain was grown for 24 hours at 28 ºC in 10 ml of Luria-Bertani (LB) liquid medium, harvested
by centrifugation at 4 ºC and resuspended in 25 ml of ultrapure H2O. After being washed three
times with water, cells were resuspended in 2 ml of 10% ice cold glycerol (J.T. Baker® Avantor™, Center Valley, PA). Aliquots (200 µl) of cells were electroporated (2.5 kV; 25 mA;
25 µF and 400 Ω) with 0.165 ng plasmid. After transformation, 1 ml LB broth was added,
incubated for 1 hour at 28 ºC, plated on LB medium supplemented with tetracycline (50 µg/ml)
and grown at 28 ºC for 48 hours. Colonies emitting GFP fluorescence under UV were purified
and used for later study.
‘Sedona’ onions were grown as described in Experiment 1. Plants were arranged in a
randomized complete block design with 4 treatments and 3 replications. The four treatments
included: (1) an undamaged control, (2) thrips injury alone, (3) P. ananatis alone and (4) thrips

38

injury in combination with P. ananatis inoculation. Plants receiving thrips were infested with 12
adult thrips/plant.
Eight days after thrips infestation, all treatments were inoculated with the GFP strain of
P. ananatis (~1 × 108 CFU/ml) using a 250 ml spray bottle until leaves were fully covered and
started to drip. Plants were maintained as described in Experiment 1.
Leaf tissue from each plant was analyzed for the GFP-tagged P. ananatis strain in order
to compare the bacterial density on leaf tissue colonized by thrips to other non-symptomatic leaf
tissue. Three leaf pieces were excised from each plant (n = 36). Areas where thrips feeding
injury was present were chosen in those treatments which received thrips (treatments 2 and 4),
symptomatic areas were chosen in treatments inoculated with bacteria (treatments 3 and 4) and
random areas were chosen in the treatment which received neither (treatment 1). Leaf tissue was
excised (1.0 cm2) and surface sterilized as described in Experiment 1. The resulting suspension
was diluted ten-fold, twice, and 50 µl of each concentration of the dilution series was streaked
onto LB medium supplemented with tetracycline (25 µg/ml) and incubated at 28 C for 48 hours.
The number of colonies with the GFP fluorescence were identified using UV light, counted on
each plate containing the lowest dilution and multiplied by 100 in order to calculate total
bacterial density. Mean bacterial density (colony forming units: CFU) per cm2 of leaf tissue was
compared between treatments using a t-test. The number of CFUs was log transformed to meet
normality requirements of a t-test.
For microscopy, onion plants were removed from pots 8 days after inoculation and
washed under running tap water. Leaf tissue (1.0 cm2) was excised and mounted on a microscope
slide. Areas where thrips feeding injury was present were chosen in those treatments which
received thrips (treatments 2 and 4), symptomatic areas were chosen in treatments inoculated

39

with bacteria (treatments 3 and 4) and random areas were chosen in the treatment which received
neither (treatment 1). Fluorescence microscopy was carried out on a Nikon Eclipse E800
epifluorescence microscope (Nikon Corporation, Tokyo, Japan) with the Nikon B – 2A
fluorescence filter (Nikon Corporation) for GFP analysis. GFP-tagged bacterial cells were
excited using a 490 nm filter and the images were captured with a Samsung Galaxy S6 Active
camera (Samsung Group, Seoul, South Korea).

Results
Sequencing of the 16S rDNA indicated that P. ananatis was present in 100% of the
treatments that were inoculated with the bacteria. None of the untreated or thrips only treatments
were positive. The most similar GenBank sequence results for the bacterial isolates were P.
ananatis (96 – 98% match, accession KT957000.1).

Experiment 1. The number of leaves per plant and total leaf area before manipulation were
statistically similar between treatments (leaf number: F = 0.95, df = 5, 70 p = 0.45; leaf area: F =
0.54, df = 5, 70 p = 0.75). No statistical differences were found among the treatments at the end
of the experiment in the number of leaves per plant, total leaf area and plant growth (leaf
number: F = 0.68, df = 5, 70, p = 0.64; leaf area: F = 1.14, df = 5, 70, p = 0.35; growth: F = 1.35,
df = 5, 70, p = 0.25; Table 3.1). All treatments inoculated with P. ananatis had significantly
more bacterial leaf blight symptoms than treatments which did not receive bacteria (t > 3.39, df =
5, p < 0.01; Table 3.1). The percentage of tissue damaged by thrips was significantly higher in
treatments with thrips alone (treatment 2; numbers correspond to treatment numbers in Table
3.1) and thrips with P. ananatis (treatment 4) compared to all other treatments (t > 4.60, df = 5 p

40

< 0.01; Table 3.1). Significantly more thrips were counted on all treatments which received
thrips relative to those without thrips (t > 3.87, df = 5, p < 0.01; Table 3.1). There were
approximately three times more thrips on the treatment that received P. ananatis with thrips
simultaneously (treatment 6) compared to the other two treatments with thrips (treatments 2 and
4) (t > 8.06 df = 5, p < 0.01; Table 3.1).

41

Table 3.1. The effects of onion thrips (Thrips tabaci), bacterial leaf blight (Pantoea ananatis) and mechanical damage (five pricks
using a pin) on onions. When thrips and bacteria were present together, bacteria were either added to plants 7 days after thrips or
simultaneously. Thrips (3-4 adults/plant) were added to plants from a laboratory colony. Bacterial (1 × 108 CFU) solution was sprayed
onto plants using a hand spray bottle. Experiments were conducted for 14 days in a growth chamber at Michigan State University,
East Lansing, MI. N = 15. Numbers (1-6) in the second row correspond to treatment numbers for experiment 1 used in the text.
Damage Parameters
Number of leaves
Total leaf area, cm2 c
Growth, cm2 d
Estimated % leaf blighte
Estimated % thrips
injuryf
Thrips per plantg

Treatmentsa,b (mean ± SE)
-/- (1)

th/- (2)

6 ± 0.17a
584.67 ±
26.78a
456.56 ±
18.95a
0.00 ± 0.00b

6.06 ± 0.18a
520.62 ±
33.98a
414.79 ±
19.40a
0.00 ± 0.00b

0.00 ± 0.00b

9.30 ± 2.41a

-/pa (3)

6.13 ± 0.17a
586.56 ±
31.16a

th/pa (4)

md/pa (5)

th/pa [st] (6)

1.93 ± 0.50a

5.93 ± 0.12a
512.45 ±
44.23a
411.34 ±
35.77a
2.13 ± 0.27a

6.13 ± 0.17a
547.62 ±
23.73a
426.10 ±
14.20a
1.46 ± 0.39a

6.33 ± 0.21a
589.38 ±
38.04a
463.46 ±
20.50a
1.80 ± 0.28a

0.00 ± 0.00b

10.60 ± 2.75a

0.00 ± 0.00b

2.60 ± 0.69b

463.15 ± 19.4a

0.00 ± 0.00c
3.86 ± 0.60b
0.00 ± 0.00c
4.26 ± 0.88b
0.00 ± 0.00c
12.60 ± 1.47a
Different letters within a row indicate differences among treatments, Tukey’s HSD, α < 0.05.
b
Thrips infestation (th), P. ananatis inoculation (pa), mechanical damage (md), thrips and bacteria added at the same time [st].
c
(Gamiely et al. 1991).
d
Growth = total leaf area prior to rating – total leaf area 7 days after inoculation.
e
Estimated percentage of leaf damage from bacterial leaf blight cause by P. ananatis (Horsfall and Barratt 1945).
f
Estimated percentage of leaf damage from thrips feeding.
g
Number of thrips counted on each plant
a

42

Experiment 2. The number of leaves per plant and total leaf area before manipulation did not
differ between treatments (number of leaves: F = 0.95, df = 3, 30 p = 0.45; leaf area: F = 0.54, df
= 3, 30 p = 0.75). The number of leaves per plant, total leaf area and plant growth were similar
among treatments after experimental manipulation (leaf number: F = 1.00, df = 3, 30, p = 0.41;
leaf area: F = 2.23, df = 3, 30, p = 0.11; growth: F = 2.28, df = 3, 30, p = 0.10; Table 3.2). The
treatment that received 20 thrips exhibited the most severe blight symptoms at the end of the
experiment, with about 35% more leaf tissue necrosis than any of the other treatments (t = 5.18,
df = 3, p < 0.01; Table 3.2). The percentage of leaf tissue damaged by thrips was 12% and 15%
higher when initial infestation was 12 and 20 thrips per plant (treatments 3 and 4; numbers
correspond to treatment numbers in Table 3.2) (t > 5.16, df = 3, p < 0.01; Table 3.2),
respectively, compared to the treatments with lower numbers of thrips. The number of thrips per
plant at the end of the experiment was seven times higher than when initial thrips infestation was
20 thrips per plant (treatment 4) (t = 2.31, df = 3, p < 0.01; Table 3.2) compared to when initial
infestation was 0 or 4 thrips per plant (treatments 1 and 2). There was a significant positive
correlation, across all treatments, between the number of thrips on each plant and the percent leaf
area damaged by P. ananatis (R2 = 0.68, p < 0.01; Figure 3.1A). Thrips feeding damage
symptoms on leaves (%) and severity of leaf blight symptoms (%) were positively correlated (R2
= 0.70, p < 0.01; Figure 3.1B).

43

Table 3.2. Effects of varying numbers of onion thrips (Thrips tabaci) per plant in combination with bacterial leaf blight (Pantoea
ananatis) on onions. Thrips (0, 4, 12 or 20/plant) were added to plants from a laboratory colony. Plants were inoculated with a liquid
culture of P. ananatis (1 × 108 CFU), sprayed onto plants. Experiments were conducted for 15 days in a growth chamber at Michigan
State University, East Lansing, MI. N = 11. Numbers (1-4) in the second through fifth rows correspond to treatment numbers for
experiment 2 used in the text.
Treatmentsa,b (mean ± SE)

Damage Parameters
0 th/pa (1)

4 th/- (2)

12 th/pa (3)

20 th/pa (4)

7.73 ± 0.20a
7.18 ± 0.26a
7.36 ± 0.20a
7.36 ± 0.28a
1006.92 ± 42.78a 821.51 ± 52.79a 877.18 ± 41.09a 956.82 ± 64.90a
558.78 ± 33.90a 374.23 ± 52.57a 446.59 ± 42.00a 486.42 ± 56.65a
11.27 ± 0.54b
14.09 ± 0.80b
13.27 ± 0.96b
19.72 ± 1.59a
0.00 ± 0.00b
3.45 ± 0.53b
15.72 ± 1.65a
18.09 ± 2.61a
0.00 ± 0.00b
1.63 ± 0.53b
6.18 ± 0.67ab
12.27 ± 3.16a
a
Different letters within a row indicate differences among treatments, Tukey’s HSD, α < 0.05.
b
Thrips infestation (th), P. ananatis inoculation (pa), mechanical damage (md), added at the same time (st).
c
(Gamiely et al. 1991).
d
Growth = total leaf area prior to rating – total leaf area 7 days after inoculation.
e
Estimated percentage of leaf damage from bacterial leaf blight cause by P. ananatis (Horsfall and Barratt 1945).
f
Estimated percentage of leaf damage from thrips feeding.
g
Number of thrips counted on each plant.
Number of leaves
Total leaf area, cm2 c
Growth, cm2 d
Estimated % leaf blighte
Estimated % thrips injuryf
Thrips per plantg

44

Experiment 3. Pantoea ananatis was not recovered from any of the treatments which did not
receive bacteria (treatments 1 and 2; numbers correspond to treatment numbers in Table 3.1).
Pantoea ananatis colonies containing the GFP gene were cultured from symptomatic leaf tissue

45

on plants which received P. ananatis without thrips (treatment 3) and P. ananatis in combination
with thrips (treatment 4) with mean bacterial densities of 5.83 × 105 and 3.01 × 105 CFU,
respectively. Fluorescent colonies were also isolated from random non-symptomatic areas on
plants which received P. ananatis alone (treatment 3) and from feeding sites on plants which
received P. ananatis in combination with thrips (treatment 4) with mean bacterial densities of
1.44 × 103 and 1.58 × 105, respectively. The average bacterial density per plant did not differ
between treatments (t = 1.75, df = 34, p = 0.09). Bacterial density on leaf tissue with disease
symptoms from plants which received P. ananatis without thrips (treatment 3) compared to
symptomatic leaf tissue from plants which received P. ananatis in combination with thrips
(treatment 4) was statistically similar (t = 0.95, df = 16, p = 0.36). On plants which received both
P. ananatis and thrips, bacterial density was significantly higher in leaf tissue containing thrips
feeding damage (treatment 4) compared to areas without thrips feeding damage (treatment 3) (t =
3.32, df = 16, p < 0.01).
Pantoea ananatis was not detected in leaf tissue without bacterial application regardless
of thrips damage (Figure 3.2A, B). Fluorescence microscopy confirmed the colonization of onion
leaf tissue by the GFP strain of P. ananatis, mainly in intercellular regions of damaged cells
(Figure 3.2C). Fluorescent P. ananatis colonized thrips feeding sites, resulting in necrotic lesions
(Figure 3.2D).

46

47

Discussion
These results suggest that onion thrips feeding injury promotes the colonization of onion
leaves by P. ananatis and bacterial symptoms are more abundant with increasing thrips numbers.
Previous research demonstrated that thrips were able to acquire this pathogen after a few hours
of feeding on an infected plant and that bacteria was able to survive transstadially without
affecting the fitness of the insect host (Dutta et al., 2014, Dutta et al. 2016). Thrips are able to
spread this pathogen to new plants via stercorarial means, i.e., through their feces (Dutta et al.
2014, Dutta et al. 2016), and their feeding injury provides entry points for bacterial infection. A
similar relationship was recorded, where the feeding injury of thrips promoted the development
of purple blotch by providing alternative penetration sites for A. porri (McKenzie et al. 1993).
By utilizing fluorescence microscopy, I was able to visualize the GFP-tagged strain of P.
ananatis colonized areas of the onion leaf tissue damaged by thrips, providing evidence that
feeding sites act as disease entry points.
Onion thrips reproduce asexually through parthenogenesis and the rate of reproduction is
dependent on environmental conditions. Under hot and dry conditions, onion thrips oviposition
rates increase and generation time decreases, leading to rapid population growth (Rueda et al.
2007, Gill et al. 2015). These results demonstrate that as the number of thrips per plant and their
feeding damage increases, so do the symptoms of bacterial center rot caused by P. ananatis
(Figure 1). These results are similar to previous findings from an insecticide trial where plots of
onions receiving insecticides through the growing season had significantly less necrotic leaf
tissue compared to plots with unmanaged thrips populations (Grode et al. 2017). Therefore, it is
important for onion growers to manage onion thrips populations, particular during hot weather,
in order to minimize the development of bacterial rot in onions, and future research should focus

48

on understanding the application threshold for insecticides for reducing thrips populations along
with bacterial symptoms.
The interaction between onion thrips and P. ananatis during the early growing season is
poorly understood, and while the inoculum source of P. ananatis is currently unknown, it has
been previously identified from onion seeds (Walcott et al. 2002) and a variety of other crops
(Bruton et al. 1986, Wells et al. 1987, Bruton et al. 1991, Azad et al. 2000, Coutinho et al. 2002).
This bacterium has also been identified as a common epiphyte of various weed species (Gitaitis
et al. 2002). Onion thrips are known to overwinter on many weed species surrounding onion
fields (Smith et al. 2011) before colonizing volunteer onions in the field during the spring
(Larentzaki et al. 2007). Monitoring and managing weed species during early growing season for
both onion thrips and P. ananatis may prevent the disease from spreading to the field. A better
understanding of this interaction may provide insights for improving early-season management
of this pest complex.
These laboratory results will need to be verified under field conditions in the future as
much less is understood about P. ananatis and thrips relationships under field conditions.
Bacterial pathogens, such as P. ananatis, can be disseminated throughout the field via wind,
splashing water (Wiriyajitsomboon et al. 2014) and contaminated machinery (Gitaitis et al.
1978), therefore the relative contribution of thrips and different abiotic factors to pathogen
spread in onion fields needs to be better understood. In addition, while field research
demonstrated that onion thrips are contributing to the spread of a closely related pathogen, P.
agglomerans (Grode et al. 2017), the acquisition rates for P. ananatis and P. agglomerans are
not the same (Dutta et al. 2014), thus results with one pathogen should not be readily assigned to
the other.

49

CHAPTER 4:
Conclusions and future directions

The purpose of this project was to determine the relationship between onion thrips and
the bacterial pathogens responsible for bacterial leaf blight and bacterial center rot in onion,
Pantoea agglomerans and P. ananatis. In the second chapter of this thesis, I examined the
relationship between onion thrips and P. agglomerans in the field in 2015 and 2016 and found
positive correlations between onion thrips population densities, numbers of onion thrips positive
for P. agglomerans, and bacterial leaf blight incidence in onion fields. During the early weeks of
the 2015 and 2016 growing seasons onions symptomatic with bacterial leaf blight reached >
60%; however, only a small proportion of onion thrips screened positive for P. agglomerans (<
10%) during this time. As the season progressed, bacterial leaf blight reached 100% incidence in
the fields by late July in 2015 and early July in 2016. The number of onion thrips positive for P.
agglomerans also increased throughout both growing seasons reaching peak positive proportions
in mid-late July. Since the proportion of onion thrips carrying P. agglomerans did not increase
until several weeks following an increase in bacterial leaf blight incidence, it is unlikely that
onion thrips are introducing the bacteria into the fields. In pesticide trials conducted in 2015 and
2016, insecticides reduced both onion thrips abundance and bacterial leaf blight incidence, while
bactericides showed no effect on controlling bacterial leaf blight severity.
In the third chapter, I examined the effects of injury to onion by onion thrips and its role
in infection by P. ananatis and development of bacterial center rot symptoms under laboratory
conditions. When inoculated with a liquid culture of P. ananatis, onions with onion thrips
feeding injury had the highest percentage of necrotic leaf tissue. Also, significant positive

50

correlations were determined between the number of thrips per plant and the severity of leaf
blight symptoms as well as the percentage of feeding damage and the severity of leaf blight
symptoms. Lastly, by utilizing a GFP tagged strain of P. ananatis, I observed the colonization of
onion thrips feeding injury by P. ananatis, and compared the bacterial concentration in onions
with disease and no onion thrips to onions with disease and onion thrips. I found that the
concentration of bacteria was higher in areas of thrips feeding injury but when the total
concentration on plants was compared no significant differences were found.
Onion thrips populations fluctuate based on temperature and populations increase
drastically when temperatures rise above 25C in the field (Rueda et al. 2007). Insect data
collected during the 2015 and 2016 growing season’s show population increases in late July, a
typically hot period during Michigan summer’s, which coincided with high incidences (> 90%)
of bacterial leaf blight in sampled fields. Similarly, over the course of late July and early August,
onion thrips positive for P. agglomerans reached highest proportions, suggesting that during
optimal weather conditions onion thrips were more actively feeding and acquiring more bacteria.
It is important to note, the time when the highest proportion of onion thrips were positive for P.
agglomerans occurs only a few weeks before when onions senesce, a time when onion thrips are
of least concern and insecticide applications are often reduced. In cases where infection by P.
agglomerans and P. ananatis is severe early senesce of onions may occur (Mary Hausbeck
verbal communication) which could also correspond with a period of increased onion thrips
pressure. During the late part of the growing season, some onion thrips undergo exodus flights
and travel long distances, often field to field (Smith et al. 2015). If a high proportion of onion
thrips making exodus flights are carrying P. agglomerans, or other related bacteria, long-distance
disease spread could occur.

51

The inoculum source of P. agglomerans as well as P. ananatis is currently unknown,
however both bacteria are found in other cropping systems as both plant pathogens and epiphytes
(Serrano 1928, Bruton et al. 1986, Wells et al. 1987, Bruton et al. 1991, Azad et al. 2000,
Coutinho et al. 2002, Procópio, 2004, Medrano and Bell 2007, Morales-Valenzuela et al. 2007,
Romeiro et al. 2007, Ferreira et al. 2008, Lee et al. 2010, Yang et al. 2011). Pantoea ananatis is
also known to naturally infest onion seeds and this has been hypothesized as an inoculum source
for bacterial disease in the field (Walcott et al. 2002). It may be important to screen onion
seedlings for P. ananatis and P. agglomerans during the early growing season to determine if the
bacteria originates from seeds. The role of onion thrips as an inoculum source for P.
agglomerans and P. ananatis has not been investigated. Before colonizing volunteer onions in
the field the following spring (Larentzaki et al. 2007), onion thrips feed and overwinter on an
assortment of weed species. Similarly, bacteria such as P. ananatis are known to be a common
epiphyte of various weed species (Gitaitis et al. 2002). If an interaction between onion thrips and
Pantoea sp. is occurring in weed species before the start of the growing season, onion thrips may
be introducing these diseases to the field. It is known that P. ananatis can persist in tobacco
thrips transtadially (Dutta et al. 2016) however it has not been investigated whether these
bacteria can persist in thrips during overwintering. According to the field data, 6% of the field
caught onion thrips were positive for P. agglomerans during the first sampling period suggesting
that thrips are either acquiring the disease from another source, such as weeds, or it is persisting
in overwintering populations. Early season screening of onion thrips populations residing in
nearby weeds may provide insight into the inoculum source of these diseases.
Results from this thesis establish a field relationship between onion thrips and P.
agglomerans and an association between onion thrips feeding damage and P. ananatis

52

development. While these bacteria are similar and are often found together in the field, is
important to investigate differences between the two and their relationship with onion thrips and
other thrips species. The laboratory results which showed a correlation between onion thrips
feeding damage and severity of bacterial center rot by P. ananatis will need to be verified under
field conditions. Prior laboratory research demonstrated that onion thrips have the ability to
acquire and transmit P. ananatis and P. agglomerans (Dutta et al. 2014). Future research will
need to explore the differences between onion thrips direct and indirect association with Pantoea
sp. directly as a vector and indirectly through feeding damage in onion fields. Aside from
transmission by onion thrips, P. ananatis and P. agglomerans are known to be disseminated
throughout onion fields via contaminated machinery (Gitaitis et al. 1978), wind or splashing
water (Wiriyajitsomboon et al. 2014). Therefore, it is also important to investigate the relative
contribution of onion thrips and other abiotic factors to the spread of Pantoea sp. throughout
onion fields.
The research presented in chapter two of this thesis is the first to investigate management
strategies which control both onion thrips and bacterial leaf blight. Based on the results of the
pesticide trail, it is clear that onion thrips are contributing to the severity of bacterial leaf blight
and the only way to reduce bacterial leaf blight incidence was to reduce the thrips populations. It
is likely that the feeding damage from onion thrips is causing the increase in bacterial leaf blight
severity. This would match the results from chapter three which established a correlation
between the number of thrips per plant and the amount of necrotic leaf tissue on onion leaves.
Current management practices utilize economic action thresholds as a way to determine when an
insecticide application is necessary (Fournier et al. 1995). Future research should focus on
investigating various application thresholds for insecticides for reducing thrips populations along

53

with bacterial leaf blight and center rot symptoms. Effectively managing onion thrips
populations could reduce the risk of losses from bulb rot during long-term storage and reduce the
continued spread of pathogens to other onion fields.

54

APPENDIX

55

Record of Depositions of Voucher Specimens

The specimens listed below have been deposited in the named museum as samples of those
species or other taxa, which were used in this research. Voucher recognition labels bearing the
voucher number have been attached or included in fluid preserved specimens.
Voucher Number: 2017-02
Author and Title of thesis:
Author: Ari Grode
Title: The Relationship between Onion Thrips (Thrips tabaci) and the Bacterial Pathogens
Pantoea agglomerans and Pantoea ananatis
Museum(s) where deposited:
Albert J. Cook Arthropod Research Collection, Michigan State University (MSU)
Specimens:
Table S. 1. Voucher specimens deposited at the Albert J. Cook Arthropod Research Collection
(Michigan State University).
Family

Genus-Species

Life Stage

Quantity

Preservation

Thripidae

Thrips tabaci

Adult

10

alcohol

56

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57

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