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EFFECTS OF HELICOSPORIDIA ON MOSQUITOES
TRACY M. CONKLIN
A THESIS PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
MASTER OF SCIENCE
UNIVERSITY OF FLORIDA
Tracy M. Conklin
I thank Dr. Drion Boucias for mentoring me through my undergraduate and
graduate education. I thank Dr. James Becnel for his expert advice and encouragement. I
also thank Dr. Verena-Ulrike Lietze; I would have been lost without her kind help. I
extend special thanks to Steven Arias, Allison McGee, Natalie VanHoose, Jessica
Noling, and Heather Furlong for their technical support. I am deeply indebted to Genie
White for her suggestions and statistical expertise. I also thank James Colee and Janice
Cole for their help with the statistics. I also thank my parents and my friends for their
prayers, love, listening ears, and wake-up calls.
TABLE OF CONTENTS
A C K N O W L E D G M E N T S ................................................................................................. iii
LIST OF TABLES ............. ..... ......................... .......... ............ vi
LIST OF FIGURES ............. .. ..... ...... ........ ....... .......................... viii
ABSTRACT ........ .............. ............. ...... ...................... ix
1 INTRODUCTION ............... ................. ........... ................. ... .... 1
2 LITERA TURE REVIEW .......................................................... ..............5
H elicosporidium History .................................................. ............................. 5
Identification of Helicosporidium as an Insect Pathogenic Alga .................................7
P anthology ................................ ........................... ........... ............... 11
In fiction C y cle ................................................................. ......... 1 1
H ost R response .................................................. ....................... ........ 15
H ost Records and Ecology.......................................................... ............... 17
Helicosporidium and M osquitoes .................................................... ...... ......... 18
Biocontrol Potential .......................... ..... .................. .... ... ... 21
In V ivo Production ...................... ...... .......... ................. .... .. .....2 1
In V itro P reduction ......... ........................................................ .. .... ..... .. 22
Storage and Stability ........... ..................................................... .. .... .... .... 23
3 M ATERIALS AND M ETHOD S ........................................ ......................... 31
Preparation of Helicosporidium....................... ........ ........ 31
C yst A m plification ........................................ ................. .... .. ... 3 1
In V itro D ehiscence .......................................................................... ..... 32
H o st R a n g e .................................... ...................................................3 2
Food Concentration Bioassays ............................................................................35
H ost D evelopm ent ....................... .................. ..... .. .. ...............3 35
Statistical A n aly ses ................................................................... ...... .. .... ... 3 6
4 R E SU L T S ....................................................... 37
Cyst A m plification and D ehiscence ........................................ ....................... 37
H o st R ang e ............... ................................................................................... 37
Age Susceptibility.................. ......... .......................... 38
Food Concentration Bioassays ............................................................................39
Host Development ................................. ... ..... ........... ......... 40
5 D ISCU SSION ............. ................. ..................................................... ......51
A M E A SU R E M E N T D A TA ....................................... .............................................60
B CONSTRUCTION OF MOSQUITO BREEDERS ...............................................62
L IST O F R E F E R E N C E S ........................................................................ .....................65
B IO G R A PH IC A L SK E TCH ..................................................................... ..................70
LIST OF TABLES
2-1 Side-by-side comparison of the filamentous cell of Helicosporidium and the
polar-capsule filament of Cnidosporidia (Microsporidia) .....................................25
2-2 N natural hosts of H elicosporidium ........................................ ........................ 29
2-3 Laboratory-produced Helicosporidium infections .........................................29
2-4 Summary of bioassay methods for four major mosquito studies.............................29
2-5 The IC50 of first instar larvae as recorded for various isolates and species..............30
2-6 Cyst production in different hosts ........................................ ........................ 30
4-1 Average SD results of assays with An. quadrimaculatus and Ae. aegypti 7
days after exposure to 1 x 104 cysts/m L.. ........................................ ............... 42
4-2 Logistic regression of host range bioassay data. ..................................................43
4-3 Average SD corrected percent mortality at 7 days post-exposure for four
instars ofAe. aegypti at five dosages of Helicosporidium .......................................43
4-4 Average SD corrected percent infection of surviving larvae at 7 days post-
exposure for different instars of Ae. aegypti at five dosages of Helicosporidia.......43
4-5 Logistic regression of age susceptibility by instar. .............................................44
4-6 Mean SD percent mortality and infection in 2, 12, and 24-hour old Ae. aegypti
after exposure to SjH e .................. ........................... ... ..... ... ........ .... 44
4-7 Logistic regression of Ae. aegypti bioassay data................................. ..............44
4-8 Mean SD infection at 1, 2, and 3 weeks post-exposure ofAe. aegypti exposed
as first instars to four dosages of SjHe. ........................................ ............... 45
4-9 Logistic regression of percent infection at 1, 2, and 3 weeks post-exposure..........45
4-10 Mean SD adult male:female ratio ofAe. aegypti at 3 weeks post-exposure to 2
d o sag es of SjH e .................................................... ................ 4 7
4-11 Corrected percent mortality over time ofAe. aegypti at three dosages of SjHe
an d fou r food lev els.......... ............................................................ .. .... .... .... .. 4 8
4-12 Mean ( SD) corrected percent mortality in Aedes aegypti 7 days after exposure
to SjHe at four different food levels and three dosages of helicosporidia ..............49
4-13 Mean ( SD) infection rates in surviving Ae. aegypti 7 days after exposure to
three dosages of SjHe at four different food levels ............................. ............... 50
4-14 Mean ( SD) fixed, FITC-labeled cysts ingested per Ae. aegypti larva exposed to
1 x 105 cysts/mL at three food levels.............. ........................ ...............50
A -1 C om piled cyst m easurem ents ........................................................................ ... 60
A-2 Compiled filamentous cell measurements...................... ...................... 61
LIST OF FIGURES
2-1 Cell types and structures of Helicosporidia ........................ .................26
2-2 Initial infection events and development of the filamentous cell ...........................27
2-3 Filament development in vivo within a hemocyte. ...............................................28
2-4 Pamelloid colony formed in vitro after several media transfers. .............................28
4-1 Percent dehiscence of five cyst preparations purified January through April
2 0 0 5 ........................................................................... 4 1
4-2 Regression of percent corrected mortality and infection on percent dehiscence of
cyst preparation at tim e of exposure.. .............................................. ............... 42
4-3 Percent mortality over time of first instar Ae. aegypti exposed to two dosages of
S jH e ........................................................................... 4 6
4-4 Average control proportion of larvae, pupae, and adults of surviving Ae. aegypti
1-3 w eeks post-exposure. ............................................... .............................. 46
4-5 Average proportion of larvae, pupae, and adults of surviving Ae. aegypti at 1-3
w weeks post exposure ............. ...... ....................... .... ...... ........ .... 47
4-6 Percent mortality at 2 days post-exposure of first instar Ae. aegypti exposed to
three dosages of Helicosporidia and four food levels........................................49
B -l Finished m osquito breeder ............................................... ............................ 64
Abstract of Thesis Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Master of Science
EFFECTS OF HELICOSPORIDIA ON MOSQUITOES
Tracy M. Conklin
Chair: Drion G. Boucias
Major Department: Entomology and Nematology
The Helicosporidia are a poorly understood group of pathogens that have been
detected worldwide in diverse groups of invertebrates. Discovered in 1921, their
taxonomic position was debated until 2002, when it was determined that Helicosporidia
are algae. Related to the vertebrate pathogen Prototheca wickerhamii Tubaki & Soneda,
Helicosporidium represents the first described algal genus of invertebrate pathogens. The
Helicosporidia are unique in that they produce a characteristic cyst stage composed of
one elongate filamentous cell wrapped around three ovoid cells within a pellicle. On
ingestion by a host, the cyst dehisces, releasing the filamentous cell and ovoid cells from
the pellicle. The filamentous cell penetrates the midgut epithelium of the host and
differentiates into the vegetative cell phenotype. Vegetative stages develop in the
hemocoel via autosporulation, and eventually form cysts.
Recently, a new isolate of Helicosporidium was discovered in Florida that infects
the blackfly Simuliumjonesi Stone & Snoddy. Bioassays were conducted with this
blackfly isolate (SjHe) against Aedes aegypti L. and Anopheles quadrimaculatus (Say).
Both species were infected at 7 days post-exposure. However, exposure to SjHe caused
melanization and high early mortality in both species. There was also high variation in
infectivity and mortality in all bioassays. In vitro dehiscence rates varied from one cyst
preparation to the next, and may account for the high levels of variation. Aedes aegypti
was selected for further bioassays due to low control mortality and synchronous
development in this species. In Ae. aegypti, susceptibility decreased significantly after the
first instar and also within the first instar with age. Most infected individuals died as
fourth instar larvae; thus infected pupae and adults were rare. Infection did not
significantly increase in Ae. aegypti maintained up to 3 weeks post-exposure.
Susceptibility also decreased with increasing food availability during exposure. Food
acted as a diluent, reducing the number of cysts ingested. Time to pupation was not
significantly delayed in infected, individually reared Ae. aegypti. The presence of
melanization, high initial mortality, and the high dosages necessary to achieve infection
indicated a non-host interaction. This interaction was unexpected and contradicts the
widely-held assumption that the Helicosporidia are not host-specific.
Since the first description by Keilin (1921), invertebrate pathogens in the genus
Helicosporidium have been detected worldwide in diverse groups of invertebrates,
including several orders of insects, mites, crustaceans, oligochaete worms, and
trematodes (Sayre and Clark 1978; Purrini 1984; Avery and Undeen 1987b; Pekkarinen
1993). Until recently, their taxonomic position remained unclear. Keilin (1921)
tentatively described Helicosporidium parasiticum as a protozoan, but Weiser (1970)
proposed that the Helicosporidia were best placed among the lower fungi. Most recently,
Boucias et al. (2001) suggested that the vegetative development of Helicosporidia was
similar to the autosporogenic growth of unicellular algae. Significantly, genetic analysis
defined the genus Helicosporidium as a member of the green algal class
Trebouxiophyceae (Chlorophyta) and, as such, it represents a novel clade of invertebrate
pathogens (Boucias et al. 2001; Tartar et al. 2002, 2003; Tartar 2004; Tartar and Boucias
2004; de Koning and Keeling 2004). The trebouxiophyte green algae are generally
photosynthetic and free-living. However, the closest relatives to Helicosporidium (the
genus Prototheca) are achlorophyllous algae that have evolved a heterotrophic life style,
opportunistically infecting vertebrates.
The infectious cyst, the stage that defines the genus Helicosporidium, comprises
three ovoid cells and a coiled, elongate, filamentous cell enclosed in an outer pellicle.
The cyst dehisces when ingested by the insect host-the pellicle ruptures, releasing the
filamentous cell and the three ovoid cells. The invasive filamentous cells pass through the
midgut epithelial layer and gain ingress to the hemocoel. Within the hemocoel, the
filamentous cells differentiate into vegetative cells, which undergo autosporogenic cell
divisions (Blaske-Lietze et al. in press). Vegetative cells have been observed to replicate
within the phagocytic hemocytes and to develop extracellularly in the hemolymph
(Blaske-Lietze and Boucias 2005). After multiple, 2- to 8-cell autosporogenic cell
divisions, a portion of the hemolymph-borne vegetative cells differentiates into cysts
(Blaske-Lietze et al. in press).
Boucias et al. (2001) reported on a Helicosporidium sp. isolated from a simuliid
(Simuliumjonesi Stone & Snoddy) by J. Becnel at United States Department of
Agriculture Agricultural Research Service (USDA ARS) in Gainesville, Florida. This
represents the fourth non-culicid nematoceran host record for Helicosporidium. Five
species of mosquitoes have been found to be naturally infected by Helicosporidium.
However, of the five recorded culicid isolates, only three have been examined in detail.
Fukuda et al. (1976) found that a Helicosporidium sp. from Culex nigripalpus Theobald
was infectious to 17 species of mosquitoes and 6 species of insects. Hembree (1979,
1981) discovered Helicosporidia infecting Aedes aegypti (L.) and Culex quinquefasciatus
Say in Thailand and later evaluated the infectivity and biological control potential of this
isolate. Seif and Rifaat (2001) performed a similar study with an isolate from Culex
pipiens L. from Egypt. Bioassays with mosquitoes have also been performed with other
isolates. Fukuda (1976) also reported melanization in mosquitoes infected with a beetle
isolate. Avery and Undeen (1987b) studied the effects of a pond water isolate on three
species of mosquitoes, noting melanization and high initial mortality. The simuliid isolate
(designated SjHe), the subject of our study, has been assayed against six species of
Diptera, three species of Lepidoptera, and a weevil (Boucias et al. 2001; Blaske-Lietze et
al. in press; Conklin et al. 2005).
The objective of our study was to further characterize mosquito-Helicosporidium
interactions; particularly, those involving early mortality and the effect of larval age and
food availability on early mortality, total mortality, and infection. Early mortality
(defined as death before the production of cysts) was common in mosquitoes exposed to
SjHe and a pond water isolate of Helicosporidium (Avery and Undeen 1987b; Conklin et
al. 2005). Their studies suggested that early mortality and melanization indicated that
mosquitoes were unsuitable hosts for these two isolates. With other mosquito isolates,
mortality occurred at the larval-pupal interface. The early mortality observed by Avery
and Undeen (1987b) and Conklin et al. (2005) may have been due to septicemia
facilitated by ingress of the filamentous cell and may be mitigated by larval age or food
level during exposure. In fact, age-based resistance to infection has been reported in three
mosquito isolates ofHelicosporidium (Fukuda et al 1976; Hembree 1981; Seif and Rifaat
2001). The effect of food availability on susceptibility has never been addressed for
Helicosporidium, but food levels may influence the number of cysts ingested. Also, the
presence of plant chemicals in the diet may alter the viability of Helicosporidium in the
Finally, we also addressed the effect of Helicosporidium infection on development.
Developmental delay due to infection has been noted in lepidopterans (Blaske and
Boucias 2004), and a weevil (Conklin et al. 2005) but has not been quantified for
mosquitoes, which have a shorter larval period than these other hosts. Hembree (1981)
stated that development of infected Ae. aegypti was delayed 1 to 2 days, but did not offer
any data on host development. Conklin et al. (2005) presented head capsule
measurements of mosquitoes exposed to Helicosporidium, which indicated that
development was affected Ae. aegypti but not in An. quadrimaculatus. Data gathered
from our experiments provide much-needed insight into the effect of Helicosporidium on
mosquitoes and early mortality in mosquitoes exposed to Helicosporidium.
Helicosporidium was discovered in 1921 by D. Keilin parasitizing larvae of a
ceratopogonid, Dasyhelea obscura (Winn.). Keilin (1921) described this new pathogen in
great detail using fixed smears and sections. His hypothesized life cycle of
Helicosporidium parasiticum included dehiscence of the spore by the filament, invasion
of the host by the ingested "sporozoites," vegetative replication in the host hemocoel, and
reformation of cysts (which later released their filaments and sporozoites after successive
drying and wetting of the dead insect).
Keilin (1921) admitted that the question of the systematic position of
Helicosporidium was difficult, ruling out its easy classification into the Cnidiosporidia
(Sporozoa), Haplosporidia, Serumsporidia, and Mycetozoa. Keilin (1921) compared the
polar capsule of Microsporidia and the spiral filament of Helicosporidium (Figure 2-1),
concluding that the similarities between these two groups were superficial. The
Haplosporidia, he argued, have a plasmodium stage and cysts surrounding the spores,
both of which never appear in Helicosporidium. Also, the spores of the Haplosporidia are
unicellular, unlike the four heterogeneous cells found in Helicosporidium. In the
Serumsporidia, Keilin (1921) noted similarities to a group of parasites infecting
Crustacea, but considered the descriptions of these species to be incomplete, making it
impossible to judge the relationship between the Serumsporidia and Helicosporidium.
Keilin (1921) saw no similarities between Helicosporidium and the Mycetozoa (slime
molds). Unlike Helicosporidium, the Mycetozoa have plasmodium and flagellate stages
and Helicosporidium has more complicated spores than the Mycetozoa. In Keilin's final
evaluation of Helicosporidium, he stated that this pathogen was a new type of Protist that
could be temporarily included in the Sporozoa. Later, Helicosporidium was provisionally
placed in its own order (Helicosporidia) within the Cnidiosporidia (Kudo 1931).
In the 1970s, there was a resurgence of interest in Helicosporidia as new isolates
were discovered in Lepidoptera, Coleoptera, and mosquitoes. Based on observations of
Keilin's original materials and an infected Hepialid caterpillar from Argentina, Weiser
(1968, 1970) suggested that Helicosporidium be moved from the Protozoa to the
primitive Ascomycetes of the family Saccharomycetaceae and subfamily
Nematosporidiae. He argued that the structure of the vegetative stages of
Helicosporidium did not resemble any of the protozoa, but he noted that the conservation
of cell shape after separation of the daughter cells in Helicosporidium was similar that of
to plants. Weiser (1970) contended that the filament of Helicosporidium was homologous
to the needle-shaped ascospores of Monosporella unicuspidata, also described by Keilin
(1920) in Dasyhelea obscure. Weiser (1970) also noted that Helicosporidium, like a
typical fungal pathogen, caused lysis of host tissues. In the Hepialid he examined,
however, the infection was limited to a cuticular wound, and it did not appear to spread
throughout the host as in Keilin's Dasyhelea larvae. Instead, tumor-like cysts with a
fibrous, multilayered lining enclosed the helicosporidial cells. No cells were found free in
the hemolymph, and many of the tumor-like cysts contained melanized cells. Weiser
hypothesized that the caterpillar was opportunistically infected through a wound, while
Dasyhelea larvae, bathed in tree wound fluid containing Helicosporidia, could be infected
along the entire body, resulting in systemic infection. The trans-cuticular infection route
was also similar to that of most entomopathogenic fungi. Soon, the first bioassays with
Helicosporidium were carried out (Kellen and Lindegren 1974), and the first electron
micrographs of Helicosporidium were produced (Lindegren and Hoffman 1976). These
studies showed that Helicosporidium could be readily transmitted per os, and that, unlike
the majority of Ascomycetes, the vegetative stages of Helicosporidium underwent mitotic
division in the nucleus and contained well-defined Golgi bodies. These characteristics
aligned Helicosporidium more closely with the Protozoa. After these studies,
Helicosporidium remained incertae sedis for more than 25 years.
Identification of Helicosporidium as an Insect Pathogenic Alga
The diagnostic feature of Helicosporidium is the cyst stage (Figure 2-1 A, B). The
cyst is round to discoid, with the appearance of a ridged biscuit (Boucias et al 2001)
measuring 4-5 [tm in width. Cyst measurements differ considerably from one isolate to
the next (Table A-i) and may be indicative of species differences. Avery and Undeen
(1987b) found that the average size of the cyst of a pond water isolate changed
significantly after passage through a heterologous host, ranging from from
5.58 0.03 [tm to 5.20 + 0.07 [tm after the first passage. The change in cyst size
presumably indicated adaptation to a new host. The cyst is composed of four cells in a
multilaminate pellicle. The pellicle is thicker in the lateral region, peripheral to the
filamentous cell (75-93 nm), than it is in the dorsal/ventral regions opposite the ovoid
cells (50-53 nm) (Blaske-Lietze et al. in press). Three ovoid cells are stacked together
along the central axis of the cell. These ovoid cells, originally referred to as
"sporozoites", each possess a peripheral nucleus enclosing a granular cytoplasm (Boucias
et al. 2001). The filamentous cell is wound around the ovoid cells, making 3 to 4 turns.
Upon physical or chemical stimulation, the cyst pellicle ruptures in the thinner,
dorsal/ventral region, releasing the filamentous cell and ovoid cells. The free filamentous
cell is highly resistant, either retaining a spiral conformation or straightening into a
needle-like shape tapered at both ends (Figure 2-1 C). The length of the filamentous cell
varies with the isolate, measuring 37 65 atm in length by 1 atm in diameter (Table A-2).
Filamentous cells are covered in short projections or barbs (340 + 60 nm), oriented in one
direction. Vegetative cells (2-4 atm) are non-motile and also have a resistant pellicle with
a textured outer surface (Figure 2-1 D, E). During autosporulation, the daughter cells are
formed within the pellicle of the mother cell. Each pellicle contains 1, 2, 4, or 8 daughter
cells (Boucias et al. 2001). The vegetative cells are uninucleate with elongate
mitochondria and well-developed Golgi bodies (Lindegren and Hoffman 1976; Blaske-
Lietze et al. in press). Once the daughter cells are released, the shell-like empty pellicle
persists (Figure 2-1 F, G) and is diagnostic of active vegetative replication (Boucias et al.
Helicosporidium shares some morphological characteristics with algae. Vegetative
replication by autosporulation is prevalent in green microalgae of the Chlorophyta,
including the achlorophyllous genus Prototheca (Bold and Wynne 1978). The Prototheca
were initially described as new organisms with homology to both yeast-like fungi and
green algae (Kruger 1894). Nadakavukaren and McCracken (1973) demonstrated the
presence of double-membraned starch storage granules (diagnostic of a plastid structure)
in Prototheca zopfii Kruger. While a plastid structure has not been localized in
Helicosporidium, the presence of a degenerate plastid is supported by molecular analysis
(Tartar and Boucias 2004; de Koning and Keeling 2004. Ultrastructural studies show that
both Helicosporidium and Prototheca contain well-developed Golgi bodies and elongate,
peripheral mitochondria (Lindegren and Hoffman 1976; Nadakavukaren and McCracken
1973). Significantly, Helicosporidium produces a specialized, diagnostic cyst stage,
which is not seen among the Prototheca. Another important difference between the two
parasites is that Prototheca only infect vertebrate hosts, while Helicosporidium is
restricted to invertebrates.
There is extensive molecular evidence for the classification of Helicosporidium as
a green alga and the relationship between Prototheca and Helicosporidium. Five genomic
sequences have been amplified and sequenced: 18S, 28S, ITS1-5.8S-ITS2, actin and P-
tubulin. All trees constructed with these sequences placed Helicosporidium with the
green algae of the class Trebouxiophyceae (Tartar et al. 2002; Tartar 2004). An EST
library has also been constructed, revealing a high number of unique sequences with no
homology to any known sequences. Using this EST library, 98% of ribosomal protein
sequences supported the green algal origin of Helicosporidium (de Koning et al. 2005).
The mitochondrial cox3 gene has also been sequenced in Helicosporidium, again
demonstrating homology to green algae (Tartar 2004). For example, a Blast N search of
the Helicosporidium cox3 open reading frame gave E values of 5e-25 for Prototheca
wickerhamii Tubaki & Soneda and 4e-13 for Nephroselmis olivacea Stein. The 16S
ribosomal DNA sequence has been obtained from a remnant plastid in Helicosporidium,
with significant homology to that of the Protothecan plastid. The plastid genome of
Helicosporidium has been further characterized by amplification of a series of highly
conserved genes known as the str-cluster. The arrangement of genes in this cluster
indicates that Helicosporidium is closely-related to P. wickerhamii, but is more derived
than its close relative, having a re-organized, highly reduced genome (Tartar et al. 2004).
Finally, the complete plastid genome of Helicosporidium has been sequenced by de
Koning and Keeling (in press), confirming the highly reduced structure of the genome.
The plastid genome of Helicosporidium is highly structured, only 37.5 kb, with very little
non-coding DNA, no inverted repeats, encoding only the minimal number of tRNAs
There is also evidence that the plastid genome of Helicosporidium is functional.
Several cDNA sequences for plastid-targeted genes have been identified in the EST
library, suggesting that the plastid has retained functions for fatty acid metabolism,
tetrapyrrole, isoprenoid, and amino acid biosynthesis. The plastid also appears to have a
high reducing potential due to the presence of ferredoxin (de Koning and Keeling 2004).
Many of these roles are shared by the apicoplast of Plasmodium, but the metabolic
diversity of the Helicosporidium plastid exceeds that of Plasmodium. The evolution of
the functional plastid of Helicosporidium may be the result of necessity, as the cyst must
subsist in the environment before it is ingested by a host, or the plastid's metabolic
diversity may be a relic of a more recent autotrophic ancestor (de Koning and Keeling
2004). Recently, Borza et al. (2005) characterized the plastid-targeted proteins in
P. wickerhamii, revealing an even greater metabolic diversity than Helicosporidium,
including carbohydrate metabolism and purine biosynthesis. This finding supports the
hypothesis that plastid reduction is continuous along a parasitism gradient from the
opportunistic parasites such as P. wickerhamii to the obligate parasites like Plasmodium
spp. Helicosporidium appears to lie between these two extremes, as an obligate parasite
with a free infectious stage with the possibility of a facultative cycle under certain
conditions (Boucias et al. 2001) and a moderately reduced plastid.
Although molecular analyses indicate a relationship between Helicosporidium and
algae, the exact relationship between Helicosporidium, Prototheca and other non-
photosynthetic algae remains unclear. Recent phylogenetic analyses of the genus
Prototheca place Helicosporidium with P. wickerhamii and Auxenochlorella
pI /Ith ,'L id, ie/' (Kruger), basal to other Prototheca species (Prototheca moriformis
Kruger, Prototheca stagnora (Cooke), Prototheca ulmea Pore, P. zopfii) (Ueno et al.
The Helicosporidia are unique in their biology. There is no known organism with a
similar cyst stage composed of one long filamentous cell coiled around three ovoid cells
within a pellicle. When this cyst is ingested, it dehisces, releasing the filamentous cell.
Dehiscence is probably a result of a combination of protease activity, and pH. Many of
the known host insects of Helicosporidium are herbivores with very basic gut pH levels
(pH 9-12), and lepidopteran gut extracts induce dehiscence (Boucias et al. 2001).
However, proteases alone do not induce dehiscence in vitro, except with pre-treatment
with the membrane permeability enhancer dimethyl sulfoxide (Blaske-Lietze et al. in
press). Mechanical pressure also induces dehiscence (Boucias et al. 2001). High gut pH
may be involved in dehiscence. Since dehiscence is necessary to achieve infection, the
cyst's ability to dehisce is a direct indication of viability.
After dehiscence, the filament and the three ovoid cells or sporoplasms are released
into the host gut. Initially, the sporoplasms were thought to be infective, while the
filamentous cell was a unique means of opening the cyst pellicle (Keilin 1921). It is now
apparent that the filament itself is the infective cell that passes the midgut epithelium to
initiate infection. In fact, the sporoplasms deteriorate in the gut (Boucias et al. 2001;
Blaske-Lietze and Boucias 2005). Hypothetically, the passage of the midgut epithelium is
aided by anchoring barbs that have been observed on the surface of the filamentous cell
under SEM (Boucias et al. 2001). However, orientation of the barbs in the gut of
lepidopteran larvae appears to indicate otherwise. Barbs are observed pointing toward the
gut lumen on the lumen side (Figure 2-2 A), but point away from the gut once through
the basal membrane (Figure 2-2 B inset) (Blaske-Lietze and Boucias 2005). Convergent
evolution has produced similar ingress mechanisms in fungi and microsporidia. The polar
filament of microsporidia is adapted to inject the parasite's cytoplasm into a susceptible
midgut cell (Bigliardi and Sacchi 2001). Oomycete Haptoglossa erumpens likewise
makes use of a specialized "gun cell" to inject the cell protoplasm into cells of its
nematode host (Glockling and Beakes 2002). In addition to these intracellular parasites,
extracellular parasites are also known to have needle-like penetration mechanisms. For
example, the pathogenic yeast Metschinikowia bicuspidata var. australis, closely related
to Monosporella unicuspidata described by Keilin (1920), has a club-shaped ascus which
liberates its two needle-shaped ascospores upon digestion with snail gut juice or
mechanical pressure, penetrating the host gut wall (Lachance et al. 1976).
Once across the midgut, the filamentous cell divides and releases the first
vegetative cells. This stage of infection is transitory, but the filamentous cell has been
observed emerging from the basal side of the midgut 4-24 hours post-exposure (Figure 2-
2 B). While these filaments appear to pass through the midgut completely, filamentous
cells are not observed to be freely circulating in the hemolymph. Instead, early vegetative
stages appear in the hemolymph and hemocytes 48 hours post-exposure (Blaske-Lietze
and Boucias 2005). The filamentous cell pellicle can be observed inside hemocytes with
early vegetative stages (Figure 2-3). Thus, the host hemocytes appear to be the site of
initial replication. The process of filament to vegetative cell transformation has also been
observed by in vitro studies (Blaske-Lietze et al. in press). In vitro, within 24 hours, the
cytoplasm and nucleus of the filament migrate to one end, causing a swelling and
shortening of the filament. Nuclear division followed by cytoplasmic division produces
4 daughter cells within the pellicle of the filament. At 48 hours, the filamentous cell is
observed to rupture along the horizontal axis of the pellicle, releasing the four elongate
cells (Figure 2-2 C), which then divide into eight oval to spherical vegetative cells. These
vegetative cells undergo autosporulation, colonizing the hemocoel of the host.
During autosporulation, nuclear division followed by cytokinesis, and formation of
daughter cell pellicles all take place within the mother cell pellicle. Helicosporidium
produces two, four, or eight daughter cells per mother cell. Secretion of new pellicles and
the shedding of the mother pellicle can occur at two, four, and eight-cell stages, but also
appears to occur often in single-celled vegetative stages, though the significance of
shedding the pellicle at this stage is unknown (Blaske-Lietze et al. in press).
Autosporulation has been observed to occur both intracellularly and in the hemocoel.
Autosporulation continues for many cycles. In vitro cultures experience a lag phase of
1.5 days and a log growth phase between 1.5 and 6 days, reaching the stationary phase at
7 days with a density of 1.27 x 105 cells/tL. The doubling time of in vitro cultures during
the exponential phase is eight hours. Likewise, in vivo, the doubling time in the
hemolymph is seven hours (Blaske-Lietze et al. in press).
Several days after infection, cysts begin to form in the host. The timing of cyst
formation may be related to the development of the host, as cyst formation has been
observed to occur more slowly in long-lived hosts (Conklin et al. 2005). The stimulus to
produce cysts in the host is unknown. Cyst formation does not occur in vitro, but
injection of in vitro produced vegetative cells consistently results in cyst formation. High
cell densities and media depletion do not appear to be important in cyst differentiation.
The death of the host insect also does not appear to affect cyst differentiation. It is
thought that the cyst is derived from the 4-cell vegetative stages which are produced at a
genetically determined rate, since the proportion of cysts produced from in vitro cultures
when injected into a host corresponds to the proportion of 4-cell vegetative stages present
in the culture (Blaske-Lietze et al. in press). Rather than forming cysts, adherent,
multicellular clusters of cells known as pamelloid colonies form during long-term in vitro
culture (Figure 2-4). These colonies reflect an inability of the cells to separate during
autosporulation, and have been observed in other green algae (Blaske-Lietze et al. in
press). Once formed, presumably the cysts are released into the environment to infect the
next generation of hosts. The method of cyst dispersal is still unknown. Cysts appear to
be retained in the insect after death (Blaske-Lietze et al. in press). Vertical transmission is
possible in lepidopteran hosts at a low frequency (Blaske and Boucias 2004). However,
vertical transmission appears to be incidental, as Helicosporidia have not been observed
invading ovaries (Blaske and Boucias 2004). Fukuda et al. (1976) found no vertical
transmission in Culex salinarius Coquillett.
The host response to Helicosporidium infection is variable. Gross signs of
infection such as slow movement and severely retarded growth have been observed in a
few hosts (Keilin 1921; Hembree 1979; Conklin et al. 2005). The primary sign of
infection is milky white hemolymph (Boucias et al. 2001; Sayre and Clark 1979; Conklin
et al. 2005). In lepidopteran hosts, infected larvae often molt into deformed pupae or
adults (Blaske and Boucias 2004).
Delayed development appears to occur in some hosts (Hembree 1981; Blaske and
Boucias 2004; Conklin et al. 2005). Hembree (1981) reported that infected Ae. aegypti
larvae pupated 1-2 days later than uninfected larvae and that pupation was lengthened.
Quantitative data on this observed delay in development was lacking, however. Blaske
and Boucias (2004) found that developmental delay varied among susceptible noctuid
species. Pupal weight was reduced in infected H. zea, but not in Tricoplusia ni (Hubner)
or Spodoptera exigua (Hiibner). Pupation was delayed in H. zea and S. exigua, but not in
T. ni. Eclosion was delayed in S. exigua, but not in H. zea or T ni. However, a high
proportion of infected adults and pupae of all species were malformed and adult
longevity of infected individuals was reduced for all species (Blaske and Boucias 2004).
Conklin et al. (2005) reported reduction in larval weight gain in infected Diaprepes
Helicosporidium, for the most part, appears to be able to effectively evade internal
host defenses. The filamentous cell is apparently attacked and phagocytosed during initial
infection events, but resists lysis within the phagosome, producing daughter cells that are
eventually released from the hemocytes (Blaske-Lietze and Boucias 2005). Boucias et al.
(2001) found that circulating hemocytes did not attack vegetative cells. Unlike
entomopathogenic fungi that reduce the number of circulating hemocytes or impede
phagocytosis, the hemocytes appear to be normal in both appearance and number in
Helicosporidia-infected hosts (Blaske-Lietze et al. in press; Boucias et al. 2001). While
the majority of Helicosporidium-host interactions have shown the remarkable stealth of
Helicosporidium, there are four reports in the literature of encapsulation, nodulation, or
melanization in response to infection. Weiser (1970) described the formation of
multicellular sheaths surrounding Helicosporidium that appeared to originate from a
cuticular wound. The cuticle had formed a callus over the wound, enclosing some
vegetative stages and cysts. Around the wound, the masses of helicosporidial cells were
encapsulated by layers of cells originating in the hypodermis with no hemocytes evident.
Helicosporidial cells were melanized in most areas. In addition to the cells encapsulated
near the cuticular wound, helicosporidial cells were found encapsulated by lymphocytes
in other areas of the host. Likewise, Boucias et al. (2001) found that Galleria mellonella
injected with purified cysts phagocytosed the cysts formed hemocytic granulomas,
melanized hemocytes attached to various tissues. Within these granulomas, vegetative
cells continued to multiply, and invaded host tissues, but no freely-circulating vegetative
cells were observed. Fukuda et al. (1976) found that a Helicosporidium isolate from a
beetle produced localized infections in mosquito larvae along with melanization, but a
mosquito isolate tested in the same study produced systemic infections with no
melanization. A pond-water isolate produced a similar response in mosquito larvae
(Avery and Undeen 1987b, Kim and Avery 1986). It is apparent that the origin of the
isolate may have an affect on whether or not the host will exhibit an immune response to
Typically, infection by Helicosporidium does not cause acute mortality in insect
hosts. Most infected insects live until pupation or even adulthood (Fukuda et al. 1976;
Hembree 1981; Blaske and Boucias 2004; Blaske-Lietze and Boucias 2005; Blaske-
Lietze et al. in press). The precise cause of death is unknown, though death at the larval-
pupal or pupal-adult interface may indicate a disruption in molting behavior or endocrine
system. At the time of death, the host hemocoel is full of helicosporidial cells. In
Helicoverpa zea, 9.7 x 106 cells are present in each microliter of hemolymph at 10 days
post-exposure. Such a large number of invasive cells may restrict hemolymph flow and
Early mortality has been reported in mosquito larvae. Fukuda et al. (1976) report
that first instar Cx. salinarius exposed to high dosages for a long period of time rarely
survived to the fourth instar, but did not mention the precise time at which this mortality
occurred. Avery and Undeen (1987b) report that the majority of mortality to mosquito
larvae occurred within 72 hours post-exposure, and attributed this response to septicemia.
Host Records and Ecology
Members of the genus Helicosporidium have been discovered worldwide,
parasitizing a wide range of invertebrate taxa. Helicosporidium parasiticum, the only
described species in the genus, was discovered in larvae of a ceratopogonid living in tree
wounds in Cambridge, England. There are 49 reports of natural Helicosporidium
infections in invertebrates, spanning 37 described species and five continents (Table 2-1).
Natural infections have occurred in mites, cladocerans, Coleoptera, Collembola, Diptera,
Lepidoptera, Oligochaeta, and parasitic trematodes. Most natural infections are associated
with hosts that inhabit moist environments such as tree wounds (Keilin 1921; Yaman and
Radek 2005), water (Avery and Undeen 1987a; Blaske and Boucias 2004; Boucias et al.
2001; Chapman 1967, 1974; Fukuda et al. 1976; Hembree 1979; Pekkarinen 1993; Sayre
and Clark 1978; Seif and Rifaat 2001), rotten fruit (Lindegren and Okumura 1973), and
forest soils (Purrini 1979, 1980, 1981). Unlike the Prototheca, which are often isolated as
free vegetative cells in sewage, soil, and tree wounds, vegetative cells of Helicosporidia
have never been isolated outside of a host.
In laboratory assays, various Helicosporidium isolates have been transmitted to 75
known species of mites, Coleoptera, Diptera, and Lepidoptera. (Table 2-2) Only one
assay has been performed on a vertebrate system. Helicosporidium was intubated into the
stomachs of 12 golden hamsters. Thirty days later the hamsters were examined and found
free of infection (Hembree 1981). Significantly, Helicosporidium does not grow in vitro
at 35C, and cysts have reduced infectivity when stored at 37C for more than a few hours
(Boucias et al 2001; Hembree 1981).
Helicosporidium and Mosquitoes
Mosquitoes are one of the best-studied hosts of Helicosporidium. The discovery of
Helicosporidium in a single larva of Culex territans Walker (Chapman 1967) first opened
the question of the biological control potential of Helicosporidium. While
Helicosporidium has been transmitted to 27 species in 8 genera of Culicidae in the lab,
natural infections have only been found in 5 species of mosquito: 4 Culex spp. and
Ae. aegypti. These isolates have originated from Lousiana, Thailand, and Egypt. Two
other nematoceran host records exist. The first described host of Helicosporidium was a
ceratopogonid (Keilin 1921), and Helicosporidium has also been isolated from S. jonesi
Of the 27 species of mosquito infected in the laboratory, 25 of these species were
infected by mosquito isolates, 7 by heterologous isolates, and 5 by both mosquito and
heterologous isolates. Mosquito isolates have been transmitted to 6 species of
Lepidoptera and Coleoptera. Anopheles spp. have been reported to be highly susceptible
to both a mosquito isolate (Fukuda et al. 1976) and a heterologous isolate (Avery and
Undeen 1987b). However, Culex spp. are also very susceptible to mosquito isolates
(Fukuda et al. 1976; Hembree 1981; Seif and Rifaat 2001). Predatory Toxorhynchites
splendens (Wiedemann) larvae are susceptible to Helicosporidium infection if fed
infected larvae (Hembree 1981).
Bioassay systems have varied considerably from isolate to isolate, making
comparisons between isolates difficult. The various bioassay methods for the major
mosquito bioassays are summarized in Table 2-3. Low dosages and long exposure times
probably best imitate natural infection conditions. Fukuda et al (1976) and Hembree
(1981) do not specify at what time post-exposure infection was assayed. Seif and Rifaat
(2001) reported that the time of assay was 10 days after exposure, while Avery and
Undeen (1987b) and Kim and Avery (1986) assayed for infection at adult emergence. In
most assays live vegetative stages or cysts indicated infection, but the infection criteria of
Avery and Undeen (1987b), included live and melanized helicosporidial cells.
In all mosquito bioassays, Helicosporidium acted in a dosage-dependent manner.
However, due to variations in bioassay methods, estimated ICso's varied greatly. For
example, the estimated IC5o of Cx. salinarius was greater than 9.1 x 106 cysts/mL when
measured by Fukuda et al. (1976), but was only 2.6 x 104 cysts/mL when measured by
Avery and Undeen (1987b). Hembree (1981) and Seif and Rifaat (2001) had comparable
exposure methods, resulting in similar ICso's for the two Aedes sp. they tested. The
estimated ICso's for each bioassay are summarized in Table 2-4. In addition to the
concentration of cysts in the exposure container, the time of exposure was directly related
to infection rates. Longer exposure led to higher levels of infection and mortality in all
bioassays (Fukuda et al. 1976; Avery and Undeen 1987b; Seif and Rifaat 2001). The
degree to which exposure time had an effect depended on the dosage tested and the age of
the larvae. If 24-hour old larvae were exposed for 24 hours, they would often molt into
the next instar during the exposure time, introducing another variable of physiological
age (Hembree 1981). Hembree (1981) exposed 24-hour old Ae. aegypti to
5 x 102 cysts/mL for a series of times from 1 hour to 48 hours. At this dosage, there was a
3-fold increase in infection by increasing exposure time from 1 to 48 hours. Seif and
Rifaat (2001), on the other hand, tested third instar Cx. pipiens at the same dosage, and
showed a 10-fold increase in infection from 1 hour to 48 hours. Results at higher dosages
(1 x 103 and 5 x 103 cysts/mL) were nearly identical for the two bioassays. Fukuda et al.
(1976) reported a 9-fold decrease in number of surviving larvae from 1 hour to 8 hours,
but the infection rate did not follow the same trend, remaining around 50% for exposure
times of 1, 2, and 8 hours.
Susceptibility decreased rapidly with larval age. Hembree (1981) notes that
physiological age instarr) rather than chronological age is the most important factor for
susceptibility. In his assays with Ae. aegypti exposed to 1 x 103 cysts/mL, percent
infection dropped from 63% to zero from 24-hour old larvae (first instar) to 48-hour old
larvae (second instar). Likewise Fukuda et al. (1976) found a 3-fold reduction in infection
rate in larvae 1 day to 3 days old. Seif and Rifaat (2001) reported that the IC5o of fourth
instar Cx. pipiens was 7-fold higher than the first instar larvae, and the second instar
larvae just 1.2-fold higher than the first instar larvae.
In order for a biocontrol agent to be successfully integrated into an IPM strategy, it
must be safe, easy to obtain, inexpensive, and able to survive storage and environmental
extremes. Helicosporidium appears to be safe to vertebrates due to temperature
limitations, though no systematic safety analysis has been performed on Helicosporidium.
The wide host range of Helicosporidium may be problematic, however, in systems where
non-target invertebrates are a concern. Feasibility of biological control has been assessed
by examining in vivo and in vitro production of cysts, storage, and the effects of
environmental conditions on cyst viability.
In Vivo Production
Helicosporidium has been successfully mass-produced in vivo in Cx. salinarius
(Fukuda et al. 1976), Ae. aegypti (Hembree 1981), and Cx. pipiens (Seif and Rifaat 2001)
as well as S. exigua (Hembree 1981), Carpophilus mutilatus (Erichson), Paramyelois
transitella (Walker) (Kellen and Lindegrean 1973), H. zea (Avery and Undeen 1987b;
Boucias et al. 2001; Blaske and Boucias 2004), and Spodoptera littoralis (Boisduval)
(Seif and Rifaat 2001). Methods for in vivo production have changed over time.
Mosquito hosts were invariably infected by exposure to a cyst suspension of a known
concentration. Hembree (1981) reports optimization procedures to determine the
appropriate dose, exposure time, and age of larvae to use to produce the highest number
of cysts per insect. Lepidopteran hosts, on the other hand, were infected by several
different methods. Kellen and Lindegren (1974) infected P. transitella larvae with diet
containing 2.6 x 106 cysts/g, though the methods by which these cysts were obtained and
integrated into the diet are not described. Hembree (1981) injected S. exigua larvae with
gradient-purified cyst preparations. Avery and Undeen (1987b) allowed starved H. zea
larvae to feed for 24 hours on a droplet containing cysts. Seif and Rifaat (2001) fed
S. littoralis on castor oil leaves treated with a droplet containing cysts. Purification
protocols likewise vary. Infected mosquitoes were collected and homogenized in
deionized water. In some studies, this was the extent of the cyst purification protocol
(Fukuda et al. 1967; Hembree 1981). Seif and Rifaat (2001) added a centrifugation step
into the purification protocol for Cx. pipiens. Lepidopteran hosts, having much more
tissue and cellular debris, were macerated and filtered, then cysts were purified on Ludox
or Percoll gradient centrifugation using methods similar to Undeen and Vavra (1998) for
purification of microsporidia (Avery and Undeen 1987b; Boucias et al. 2001).
The number of cysts produced in different insect hosts is summarized in Table 2-6.
Although cyst production in lepidopteran hosts has been reported to be 2-3 orders of
magnitude higher than mosquito hosts, infectivity of cysts may be reduced by
amplification in a heterologous host. For example, Hembree (1981) reported that cysts
produced in S. exigua were less infectious to Ae. aegypti than cysts produced in
Ae. aegypti. Seif and Rifaat (2001) also produced cysts in a lepidopteran host but did not
compare the infectivity of lepidopteran-produced cysts with mosquito-produced cysts.
Interestingly, Avery and Undeen (1987b) reported significant changes in cyst size after as
little as one passage through a heterologous host, indicating a rapid shift in phenotype
based on host (Avery and Undeen 1987b).
In Vitro Production
Helicosporidium is capable of growth on many kinds of media. Boucias et al.
(2001) found that vegetative growth was possible on insect tissue culture medium,
Candida liquid broth, Vogel-Bonner minimal broth, and SD broth. The only medium
tested that did not support vegetative growth was Czapek Dox broth, which only supports
growth of organisms capable of using inorganic sources of nitrogen. Cysts placed in
enriched growth media will dehisce, releasing filamentous cells which produce vegetative
cells (Boucias et al. 2001). Vegetative replication continues for many cycles in vitro,
eventually resulting in pamelloid colonies (Blaske-Lietze et al. in press). Cyst formation
has never been observed in vitro. More research is needed to understand the cues
requisite for cyst formation. If the signals can be determined, in vitro production of cysts
may be possible in the future.
Storage and Stability
Storage of Helicosporidium has been addressed several times over, but it is difficult
to compare many of these experiments due to differences in methods or time of storage.
High temperatures consistently deactivate Helicosporidium. Hembree (1981) reported
that Helicosporidium exposed to 420C for 24 hours has significantly lower infectivity,
and exposure to 500C for even 15 minutes reduced transmission at the highest
concentration to only 4%. However, cysts withstood 24C and 32C for 24 hours with no
loss of infectivity. At room temperature, storage for 10 days reduced infectivity to only
28% at a dosage of 3 x 105 cysts/mL, and after 17 days of storage at room temperature,
infectivity dropped to 6%. In vitro, vegetative stages do not replicate at 350C, and cells
exposed to 350C for 4 days are killed (Boucias et al. 2001).
Cold storage tolerance varies among Helicosporidium isolates. In general, purified
cysts retain infectivity well when stored in deionized water at 4-50C, an average
household refrigerator temperature (Hembree 1981; Seif and Rifaat 2001; Avery and
Undeen 1987b). Hembree (1981) reported that, when stored at 4-50C, cysts began to lose
infectivity around 6 weeks. Seif and Rifaat (2001), however, reported that storage at the
same temperature preserved infectivity through 6 months. Avery and Undeen (1987b)
reported that the LC5o of cysts stored for 3 months at 5C increased from 8.3 x 103 to
4.3 x 105 cysts/ml. At -150C, or household freezer temperature, Avery and Undeen
(1987b) and Seif and Rifaat (2001) reported that infectivity was retained for 6 months.
The data of Seif and Rifaat (2001) indicated that cysts frozen at -150C began to show
decline quicker than the cysts stored at 5C. Hembree (1981) also reported that cysts
frozen at -700C in cryoprotectant retained infectivity after 6 months of storage, and
storage at this temperature without cryoprotectant for 6 months destroyed infectivity. It is
possible that cryoprotectants at ultra-low temperature may be a long-term storage
The effect of dessication on cyst viability is also variable. Hembree (1981) found
that lyophilized and vacuum-dried cysts completely lost infectivity after 4 weeks at room
temperature. Avery and Undeen (1987b) air-dried cysts at 5C, and found that the LC5o of
dried cysts stored for 5 days increased from 8.3 x 103 cysts/mL to 9.2 x 104 cysts/mL,
indicating a 10-fold loss of infectivity. Seif and Rifaat (2001) found that air-dried spores
held at room temperature had lost all infectivity when assayed after 3 months of storage
Hembree (1981) performed a series of experiments evaluating the effect of
environmental conditions on Helicosporidium infectivity. One hour exposure to UV light
destroyed cyst viability. Exposure to buffer solutions with pH ranging from 10.5 to 3.0
for 24 hours at 40C had no significant affect on viability. A 5% solution of household
detergent for 24 hours at 40C had no effect on infectivity, but a 10% solution of detergent
reduced infectivity by 20-43%. Exposure to saline (NaC1) for 24 hours eliminated
viability at 1.71 M (10% NaCl by weight). At 0.85 M (5%), infectivity was reduced by
8 to 52%.
Table 2-1. Side-by-side comparison of the filamentous cell of Helicosporidium and the
polar-capsule filament of Cnidosporidia (Microsporidia) modified from Keilin
(1921). Later it would be shown that the filamentous cell does in fact unroll in
the intestine of a host, demonstrating convergent evolution of these two
pathogen ingress mechanisms.
Spiral filament ofHelicosporidium Polar-capsule filament of Cnidosporidia
(1) Filament is not enclosed in a polar (1) Filament is enclosed in a capsule of
capsule but lies free beneath the wall of which it forms a part.
(2) Filament always unrolls in the dead (2) Filament does not unroll until spores
body of its host. reach intestine of a second host.
(3) Filament unrolls slowly. (3) Filament is shot out.
(4) Filament is pointed at both ends and is (4) Filament is pointed at only one end and
wide and ribbon-like in the middle. very fine.
(5) Axial portion of filament is very (5) No chromatic axial portion, degenerated
chromatic, nucleus is well formed in nucleus upon wall of terminal capsule.
anterior third of filament.
(6) Filament is robust and very resistant in (6) Filament fine and very fragile.
Figure 2-1. Cell types and structures of Helicosporidia. A) SEM image of the cyst.
B) TEM image of the cyst. C) SEM image of a filamentous cell being released
from a cyst: note barbs on surface. D) and E) SEM and TEM images of the
vegetative cell. F) and G) SEM and TEM images of the vegetative pellicle.
(Boucias et al. 2001; Blaske-Lietze et al. in press, used with permission)
Figure 2-2. Initial infection events and development of the filamentous cell. A) Shows the
filamentous cell embedded in the midgut microvilli, B) the filamentous cells
emerging from the basal lamina of the midgut B inset: note orientation of
barbs is away from gut lumen once through the basal lamina, C) the in vitro
development of the filamentous cell, producing 4 elongate daughter cells
which divide into 8 spherical vegetative cells. (Boucias et al. 2001; Blaske-
Leitze and Boucias 2005; Blaske-Leitze et al. in press, used with permission)
Figure 2-3. Filament development in vivo within a hemocyte. Arrows indicate early
vegetative cells and remainder of filamentous cell pellicle. (Blaske-Leitze and
Boucias 2005, used with permission)
Figure 2-4. Pamelloid colony formed in vitro after several media transfers. (Blaske-Lietze
et al. in press, used with permission)
Table 2-2. Natural hosts of Helicosporidium
*2 identified speci
13 USA, Mexico,
10 England, USA,
3* Germany, USA
Kellen and Lindegren 1973; Lindegren
and Okmura 1973; Purrini 1980, 1985;
Blaske and Boucias 2004; Yaman and
Keilin 1921; Purrini 1981, 1984
Keilin 1921; Chapman 1967, 1974;
Hembree 1979; Fukuda et al. 1976;
Purrini 1980; Seif and Rifaat 2001;
Boucias et al. 2001
Purrini 1984; Avery and Undeen 1987a
Sayre and Clark 1978
Purrini 1980; Pekkarinen 1993
Table 2-3. Laboratory-produced Helicosporidium infections
Group Species References
Acarina 3 Kellen and Lindegren 1973
Coleoptera 13 Kellen and Lindegren 1973; Fukuda et al. 1976;
Conklin et al. 2005
Diptera 26 Kellen and Lindegren 1973; Sayre and Clark
1978; Hembree 1979, 1981; Fukuda et al. 1976,
1985; Kim and Avery 1986; Avery and Undeen
1987a,b; Seif and Rifaat 2001; Boucias et al.
Lepidoptera 11 Kellen and Lindegren 1973; Fukuda et al. 1976;
Kim and Avery 1986; Avery and Undeen
1987a,b; Hembree 1981; Seif and Rifaat 2001;
Boucias et al. 2001; Blaske and Boucias 2004
Table 2-4. Summary of bioassay methods for four major mosquito studies
Exposure Dose Volume Number
Authors Isolate source time (hours) (cysts/mL) (mL) of larvae
Fukuda et al. Cx. nigripalpus, 1-8 2 x 105 to 5 500-
1976 Cx. salinarius 1 x 107 1000
Hembree 1981 Ae. aegypti 24 5 x 102 to 20 100
Avery and Pond water 24 1 x 103 to 100 150
Undeen 1987b 4 x 105
Seifand Rifaat Cx. pipiens 24 5 x 102 to 100 100
2001 8 x 103
Table 2-5. The IC50 of first instar larvae as recorded for various isolates and species.
Authors Species IC5o (cysts/mL)
Fukuda et al. 1976 An. quadrimaculatus 1.7 x 10'
Cx. salinarius > 9.1 x 106
Hembree 1981 Ae. aegypti 1 x 103
Avery and Undeen 1987b An. quadrimaculatus 4.4 x 102
Ae. aegypti 2.2 x 104
Cx. salinarius 2.6 x 104
Seif and Rifaat 2001 Cx. pipiens 1.9 x 102
Cx. atennatus 5 x 102
Cx. perexiguus 5 x 102
Cs. longiareolata 1.7 x 103
Ae. caspius 1.4 x 103
Table 2-6. Cyst production in different hosts
Authors Species (cysts/individual)
Hembree 1981 Ae. aegypti 3.1 x 10'
S. exigua 2.0 x 107
Seif and Rifaat 2001 Cx. pipiens 8.1 x 105
S. littolaris 6.2 x 108
Avery and Undeen 1987b H. zea 7.4 x 108
MATERIALS AND METHODS
Preparation of Helicosporidium
The SjHe isolate of Helicosporidium from the black fly, S. jonesi was collected in
2002 and 2003 by J. Becnel at USDA ARS in Gainesville, Florida (Boucias et al. 2001)
SjHe was amplified in Helicoverpa zea (Boddie) and extracted on a continuous gradient
of Ludox HS40 (Perkin Elmer Life Sciences, Boston MA) following the protocol of
Blaske & Boucias (2004). Eggs ofH. zea were purchased from USDA, ARS, Stoneville,
MS. Neonates and larvae were provided with a wheat-germ-based, semi-synthetic diet
containing antimicrobial agents and preservatives (Shore and Hale 1965). Neonates were
hatched in groups and transferred individually to 24-well plates with diet after reaching
the second or third instar. All larvae were maintained at constant conditions (27 1 C,
70 5% RH, photoperiod of 12:12 [L:D] hours). Fourth and fifth instar larvae ofH. zea
were injected with 5 pL of cyst suspension at 2 x 105 cysts/insect. After injection, larvae
were transferred to individual diet cups and incubated two weeks as above. Infected
pupae and late-instar larvae were homogenized in 200 mL of deionized water with a few
crystals of 1-Phenyl 2-Thiourea. The homogenate was filtered twice through paper
toweling then subjected to two cycles of low-speed centrifugation (4,000 x g, 10 min).
The resulting pellet was layered on top of a Ludox gradient formed with a gradient maker
from 5% to 60% Ludox in deionized water. After high-speed centrifugation (16,000 x g)
for one hour, the cyst-containing band was collected and subjected to several cycles of
low-speed centrifugation (4,000 x g, 10 min) to remove residual gradient material. All
cyst preparations were stored in deionized water at 40C until use. The number of cysts in
each preparation was determined with a hemacytometer using the average of two counts.
In Vitro Dehiscence
Lepidopteran gut extracts were collected from H. zea following the protocol of
Boucias et al. (2001). Midguts of fourth or fifth instar H. zea were dissected out and
homogenized gently, then centrifuged at 16,000 x g for 15 minutes. The supernatant was
removed, passed through a MC centrifugal filter unit (0.45 [am, Millipore Corp., Bedford,
MA) and frozen at -200C in 100 aL aliquots. Rate of dehiscence was evaluated before
assays by thawing an aliquot of gut extract and adding 1 x 106 cysts to the gut extract and
gently mixing. After incubation at room temperature for 30 minutes, the cysts and
filaments were washed in deionized water 2 times using a microcentrifuge at 2,000 x g
for 10 minutes. The pellet was resuspended in 100 aL of deionized water and the percent
dehiscence was quantified with a hemacytometer.
Bioassays assessed the activity of SjHe against two mosquito species,
An. quadrimaculatus and Ae. aegypti. Mosquitoes were obtained from colonies
maintained at the USDA ARS in Gainesville, FL. Anopheles quadrimaculatus were
obtained as first instars, while Ae. aegypti eggs were obtained from the colony and
hatched in 10 mL deionized water with 1 mg of hatch media (finely ground alfalfa and
potbelly pig chow mixture (2:1)) under a vacuum. First instar mosquitoes were
transferred using a Pasteur pipette and counted into an enamel pan. Larvae were collected
on a fine-mesh cloth was and inverted into the bioassay container. For each bioassay,
100 first-instars were placed in a petri dish with 98 mL of deionized water amended with
a 1 mL dose of SjHe (treatment) or deionized water (control) and a 1-mL volume of 2%
alfalfa and potbelly pig chow mixture (2:1) as a nutritional source (final food
concentration 0.2 mg/mL). Larvae were incubated at constant conditions (26 1C,
photoperiod of 12:12 [L:D] hours) for 24 hours then transferred to enamel pans, and
water was added to a volume to 500 mL. Food was provided ad libitum. After 7 days, the
surviving individuals were counted in each pan and a sub-sample of 12 examined
randomly for infection under phase-contrast optics. Individuals containing live
helicosporidial cells (vegetative or cyst stage) were considered infected. For each species,
three replicates were performed at 1 x105 cysts/mL. Each replicate was accompanied by
an untreated control.
All further assays were carried out with Ae. aegypti, due to the low control
mortality and synchronous development of this species. Large groups ofAe. aegypti were
hatched and transferred in groups of 500 neonate larvae to enamel pans with 500 mL of
water. To obtain second, third, and fourth instars, larvae were reared at constant
conditions (26 1C, photoperiod of 12:12 [L:D] hours) and checked daily for
development. Food was provided ad libitum. Immediately after molting into the
appropriate stage, the larvae were transferred to large petri dishes in groups of 100 as
above. The four larval instars were treated at the following dosages. First instars:
1 x 103 to 1 x105 cysts/mL, second instars: 1 x 104 to 5 x105 cysts/mL, third instars: 1 x 104
to 1 x 106 cysts/mL, and fourth instars: 1 x 104 to 1 106 cysts/mL. An untreated control
group was included for each larval instar in each replicate. Larvae were assayed as above
at 7 days post-exposure. If larvae were held long enough for adults to emerge, larvae
were transferred to specially constructed mosquito breeders (see Appendix B) rather than
enamel pans, allowing for collection of adults. Adults were provided with cotton balls
soaked in 10% sucrose solution. When present, surviving pupae and adults were
examined for infection at 7 days post-exposure in addition to surviving larvae. The
percent infection of surviving pupae and adults was included in the percent infection.
The effect of age ofAe. aegypti within the first instar was also examined at
1 x 104 and 5 x 104 cysts/mL. Within two hours of the beginning of hatching, Ae. aegypti
larvae were counted into groups of 500 as above, placed in 500 mL of deionized water
with food, and incubated for 12-24 hours. First instar larvae two hours old were assayed
at the same time as 12 hour old and 24 hour old first instar larvae. In each assay, 100
larvae were exposed to Helicosporidia for 24 hours with 0.2 mg/mL of food as above. An
untreated control group was included for each larval age and replicate. After 7 days,
surviving larvae were counted and a random sample of 12 examined for infection.
Additional experiments were conducted to examine the possibility of latent
infections that become detectable only in pupae or adults. Groups of 100 larvae were
exposed as first instars to dosages from 5 x 102 to 1 x 104 cysts/mL as above. These were
maintained up to 3 weeks post-exposure, fed ad libitum at constant conditions (26 + 1 C,
photoperiod of 12:12 [L:D] hours). The number of larvae, pupae, and adults were counted
every 2 days. Individuals were assayed for infection at 1, 2, and 3 weeks post-exposure
using a random subsample of larvae, pupae, and adults.
Food Concentration Bioassays
The effect of food concentration during exposure to Helicosporidia was examined
in first instar Ae. aegypti. First instar larvae were counted into groups of 25 and exposed
to 1 x103, 1 x104 or 1 xl05 cysts/mL with 0.05, 0.1, 0.2 or 0.4 mg/mL of food for
24 hours at constant conditions as above. After 24 hours of exposure to food and
Helicosporidia, the larvae were transferred to enamel pans, maintained for 7 days and
assayed for infection as above. Mortality was recorded daily.
To further examine the effects of food concentration on ingestion of Helicosporidia,
cysts were fixed in 2.5% glutaraldehyde for at least 5 hours, rinsed and labeled with
Fluorescein isothiocyanate (FITC) in sodium bicarbonate buffer (pH 9.5) overnight. The
resulting fixed, labeled cysts were rinsed 3 times in deionized water in a microcentrifuge
at 2,000 x g for 10 minutes. Cysts were counted with a hemacytometer and added to
100 mL of deionized water with 25 first-instar Ae. aegypti at a dosage of
1 x105 cysts/mL. Three different food concentrations (0.1, 0.2, 0.4 mg/mL) were tested.
After 1, 2, or 4 hours of exposure at 260C, all 25 larvae were removed and rinsed three
times in deionized water. The larvae were homogenized in 0.05% SDS by sonication for
15 seconds, then the labeled cysts in the homogenate were quantified with a
hemacytometer. The concentration of cysts in the suspension was used to calculate the
average number of cysts ingested per insect.
To examine the effect of infection on development time, groups of 100 first instar
Ae. aegypti larvae were exposed 1 x 104 cysts/mL of SjHe for 24 hours as above. Larvae
were rinsed and each transferred into a well of a 24-well plate using a Pasteur pipette.
Each well contained 3 mL of water and 0.02 mg/mL of food. Larvae were held at 260C
and evaluated every day for molting, pupation, or death. Food was provided ad libitum
until pupation. Two plates of control and two plates of exposed larvae were used.
Individuals were assayed for infection after adult emergence.
Where control mortality was not necessary to statistical analyses, mortality of each
assay was corrected with the control group mortality using Abbott's formula (Abbott
1921). Statistical analyses were done with the SAS System for Windows (SAS Institute
1999). Percent infection and percent mortality data were subjected to logistic regression
using proc genmod. Ingestion of cysts, sex ratios, dehiscence data, and time to pupation
were subjected to analysis of variance by the procedure for general linear models (glm) in
balanced designs and the procedure for mixed linear models (mixed) in unbalanced
designs (Neter et al. 1990; Rao 1998; Younger 1998). The means were separated by the
least square means statement (ls means).
Cyst Amplification and Dehiscence
In vitro dehiscence of cysts of SjHe amplified from H. zea was highly variable.
Dehiscence rates collected from five selected cyst preparations amplified from January to
April 2005 varied from 34 to 70% immediately after purification. Percent cyst dehiscence
also changed during several weeks of storage at 4C (Figure 4-1), rapidly declining 1-2
weeks after purification. After this initial decline, dehiscence rates remained at a low,
constant baseline below 10%. In bioassays with Ae. aegypti, infection and mortality
increased with increasing percent dehiscence (Figure 4-2). Two of the 5 cyst preparations
subjected to bioassays had dehiscence rates above 20% (2/9/05 and 3/9/05), and produced
an average of 32 15% mortality, and an average percent infection of 53 20%. Other
cyst preparations having dehiscence rates below 10% below 10%, and produced an
average percent mortality of 17 24% produced average percent infection of 20 + 11%.
Both Ae. aegypti and An. quadrimaculatus were susceptible to infection with SjHe
(Table 4-1). Overall, susceptibility to infection and mortality was higher for
An. quadrimaculatus. Early mortality, represented by 1-day percent mortality, was higher
for An. quadrimaculatus in SjHe-treated groups (df= 1, P < 0.0001, 2 = 173.2), but
control mortality at 1 day post-exposure was also significantly higher for An.
quadrimaculatus (df = 1, P = <0.0001, x2 = 37.1). A similar trend held for 7-day
mortality (Table 4-2).
At 7 days post-exposure, the hemolymph of infected mosquito larvae was filled
with vegetative stages and cysts. Melanized helicosporidial cells were also observed in
the head and thorax region of infected mosquitoes. Infection of surviving
An. quadrimaculatus at 7 days post-exposure was higher than Ae. aegypti (df= 1, P
<0.0001, 2 = 65.9). All measurements of mortality and infection were dose-dependent.
Melanization, however, was statistically independent of dosage in both species (df= 1,
P = 0.6872, 2 = 0.16). Melanization was also independent of infection (df= 1,
P = 0.4596, x = 0.55). However, percent melanization in surviving individuals 7 days
post-exposure was significantly higher in Ae. aegypti than An. quadrimaculatus (df= 1, P
<0.001, 2 = 49.27).
There was a pronounced decrease in susceptibility with increasing larval age of Ae.
aegypti (Table 4-3, 4-4). Statistical analysis by probit to obtain LD50 values for each age
could not be carried out due to lack of consistent mortality or infection above 50% at
7 days post-exposure. However, at 7 days post-exposure to 1 x 105 cysts/mL, first instar
larvae had significantly higher mortality and infection rates than second (df= 1,
P <0.0001, 2 = 204.88) or third (df= 1, P < 0.0001, 2 = 254.15) instar larvae. Overall,
third and fourth instar larvae were the least susceptible to infection or mortality.
Age within the first instar also had an effect on susceptibility ofAe. aegypti. When
exposed at 2 hours post-hatch, Ae. aegypti exhibited higher early mortality (represented
by 3-day percent mortality) and 7-day mortality than 12-hour old larvae or 24-hour old
larvae (Table 4-6, 4-7). Susceptibility to infection was also reduced in 24-hour old larvae
when compared to 2-hour old larvae (df= 1, P < 0.0001, X2 = 17.4). Aedes aegypti larvae
exposed to 5 x 103 cysts/mL 24 hours after hatching were resistant to infection.
The percent infection of surviving Ae. aegypti larvae at 7 days post-exposure did
not differ significantly from the percent infection in larvae, pupae, or adults sampled at 2
or 3 weeks post-exposure (Table 4-8, 4-9). Infection rates were low for live pupae and
adults sampled at 3 weeks post-exposure. Of 141 total adults examined, 7 were found
infected. Of the 25 total pupae examined, only one was found infected. Mortality was
high in the first 3 days post-exposure, and again increased after the second week post-
exposure (Figure 4-3). Dead, infected larvae and pupae were recovered 7-14 days post-
exposure. The onset of pupation and adult emergence occurred at the same time for
control and SjHe-treated groups (Figure 4-4 and 4-5). However, groups treated with SjHe
had a higher proportion of adults at 3 weeks post-exposure when compared with
untreated controls, due to the death of larvae and pupae in SjHe-treated groups. There
were more than twice as many adult males as females in control groups at 3 weeks post-
exposure. Sex ratios in groups treated with SjHe were slightly weighted toward females,
but not significantly (df= 4, P = 0.4711, F = 0.99) (Table 4-10).
Food Concentration Bioassays
Increased food availability during exposure to SjHe decreased mortality and
infection in Ae. aegypti larvae. Total mortality increased 1-3 days post-exposure, leveling
off after 3 days post-exposure such that, in most food and dose combinations, 3-day
mortality accounted for 50-90% of total mortality at 7 days post-exposure (Table 4-11).
At two days post-exposure, mortality was significantly affected by both dosage and food
level (df= 6, P = 0.0025, F = 4.63) (Figure 4-6). The relationship between food and
mortality at 7 days post-exposure was significant at the 1 x 103 dosage (df= 3,
P = 0.0145, F = 3.98) (Table 4-12). The effect of food level on infection at seven days
was only statistically analyzed for the 1 x 103 dosage due to high mortality (-100%) in
the other treatments. At 1 x 103 cysts/mL dosage, increased food levels decreased
infection (df= 3, P = 0.0278, F= 5.19) (Table 4-13).
The number of fixed, FITC-labeled cysts ingested by first instar Ae. aegypti
decreased with increasing food availability (df= 2, P = 0.0061, F = 6.31) (Table 4-14).
The number of cysts ingested per insect was not significantly different for 1, 2, or 4 hours
post-exposure (df= 2, P = 0.5719, F = 0.57). However, ingestion rates were highly
Infection with SjHe had no effect on time to pupation for individually-reared Ae.
aegypti. Of the 48 exposed larvae transferred into individual wells, 33 survived to
pupation. The infection rate in these pupae was 63%. Time to pupation was not
significantly delayed in infected individuals when compared with uninfected individuals
(df= 4, P = 0.9010, F = 0.26). Average time to pupation in infected larvae was
9.4 + 1.7 days, while the average time to pupation for control larvae was 8.6 + 1.5 days.
Storage Time (Weeks)
Figure 4-1. Percent dehiscence of five cyst preparations purified January through April
2005. After purification, cyst preparations were stored at 4C. Cyst
preparations were assayed for dehiscence immediately after purification
(0 weeks) and each week after purification. Cysts were treated with
lepidopteran gut extracts, incubated at room temperature for 30 minutes, then
the percent of filamentous cells released was counted with a hemacytometer to
obtain percent dehiscence. Data provided by V. Lietze.
I I II
70 1/4/05 Infection
y = 1.3357x + 14.108 U 2/9/05 Mortality
.60 2 = 0.59 2/9/05 Infection
50 )K 3/9/05 Mortality
S. *0 3/9/05 Infection
o 40. ** + 3/30/05 Mortality
S. 3/30/05 Infection
S30 -- 4/20/05 Mortality
o 4/20/05 Infection
20 -- Linear (Mortality trendline)
y = 0.7356x + 12.615 Linear (Infection trendline)
+ R2 = 0.2001
0 5 10 15 20 25 30 35
Figure 4-2. Regression of percent corrected mortality and infection on percent dehiscence
of cyst preparation at time of exposure. Each point represents a single
bioassay with one of the five isolates purified from Jaunuary to April 2005.
Groups of 100 first instar Ae. aegypti were exposed for 24 hours to
1 x 104 cysts/mL. Percent mortality was obtained at 7 days post exposure,
using Abbott's correction for control mortality. Percent infection was obtained
by examining a subsample of surviving larvae at 7 days post-exposure.
Table 4-1. Average + SD results of assays with An. quadrimaculatus and Ae. aegypti
7 days after exposure to 1 x 104 cysts/mL. Infection and melanization were
measured in a subsample of surviving larvae 7 days after exposure.
Day 1 Day 7
Dose D 7 Percent Percent
Species .Percent Percent
species (cysts/mL) Percent Percent Infection Melanization
15+30 38+28 0+0 0+0
An. quadrimaculatus Control 5 a (3 36) b (3, 36)
(5) (7) (3, 36) (3,36)
S03 34 25 78 10 69 43 (4, 18 27
(4) (4) 43) (4,43)
4 34+33 72 13 84+20 8+14
(6) (5) (10, 111) (10, 111)
Ae. u,.i Control 12 5+4 0+0 0+0
(5) (6) (3,36) (3,36)
1 1 11 + 13 (3, 44 13
(3) 1 ( 36) (3,36)
1 04 1 ) 57 19 33 30 (6, 57 14
(8) 72) (6,72)
a Number of replicates, each consisting of 100 larvae.
b Total number of individuals examined across all replicates appears in parentheses after
number of replicates.
Table 4-2. Logistic regression of host range bioassay data (proc genmod).
Response Variable df X2 P
Day 1 Mortality Dose (1 x 103 vs 1 x 104) 1 6.1 0.0136
Species (Control) 1 37.1 <0.0001
Species (SjHe) 1 173.2 <0.0001
Day 7 Mortality Dose (1 x 103 vs 1 x 104) 1 60.5 <0.0001
Species (Control) 1 148.5 <0.0001
Species (SjHe) 1 214.7 <0.0001
Percent Infection Dose 1 14.3 0.0002
Species 1 65.9 <0.0001
PerenDose 1 0.16 0.6872
Species 1 49.27 <0.0001
Infection 1 0.55 0.4596
Table 4-3. Average SD corrected percent mortality at 7 days post-exposure for four
instars ofAe. aegypti at five dosages of Helicosporidium.
Instar 1 x 103 1 x 104 1 x10 1 x 106
1 7(1)a 23 23 (4) 83 24 (3) nt
2 1 (1) 5 4 (3) 19 22 (3) nt
3 2 (1) 3 (1) 4 5 (4) 12 (1)
4 ntb nt 0 (1) 0 (3)
a Number of replicates appear in parentheses. Each replicate consisted of 100 larvae.
Abbott's correction for control mortality was used.
b Not tested
Table 4-4. Average SD corrected percent infection of surviving larvae at 7 days post-
exposure for different instars of Ae. aegypti at five dosages of Helicosporidia.
Instar 1 x 103 1 x 104 1 x 105 1 x 106
1 25(1,12)' 42 30 (4, 48) 75 25 (3, 17) nt
2 0(1,12) 6 10(3, 36) 6 5 (3, 36) nt
3 0(1,12) 0(1, 12) 2 4 (4,48) 25(1, 12)
4 ntb nt 8 (1, 12) 14 17 (3, 36)
a The number of replicates appears in parentheses, followed by the total number of larvae
examined across all replicates. Each replicate consisted of 100 larvae.
b Not tested
Table 4-5. Logistic regression of age susceptibility by instar (proc genmod).
Response Variable df X2 P
Day 7 Mortality Instar (1 vs 2) 1 204.88 <0.0001
Instar (1 vs 3) 1 254.15 <0.0001
Instar (2 vs 3) 1 36.41 <0.0001
Percent Infection Instar (1 vs 2) 1 9.45 0.0021
Instar (1 vs 3) 1 21.18 <0.0001
Instar (2 vs 3) 1 9.60 0.0019
Table 4-6. Mean SD percent mortality and infection in 2, 12, and 24-hour old
Ae. aegypti after exposure to SjHe and at 5 x 103 and 1 x 104 cysts/mL.
Dose Age Day 3 Day 7 Percent
(cysts/mL) (hours post- N' Percent Percent Iec b
(cysts/mL) c c Infection
hatch) mortality mortality
5 x 103 2 3 35 26 49 34 40 13 (36)
28 + 43
80 + 15
27 + 45
10 21 (48)
25 22 (26)
29 34 (39)
19 + 34 (36)
SNumber of replicates, consisting of 100 larvae each.
b Total number of larvae examined across all replicates appears in parentheses.
c Abbott's correction for control mortality was used.
Table 4-7. Logistic regression ofAe. aegypti bioassay data for age and dose.
Day 3 Mortality
Day 7 Mortality
Age (2h vs 12h)
Age (2h vs 24h)
Age (12h vs 24h)
Age (2h vs 12h)
Age (2h vs 24h)
Age (12h vs 24h)
Age (2h vs 12h)
Age (2h vs 24h)
Age (12h vs 24h)
1 x 104
Table 4-8. Mean SD infection at 1, 2, and 3 weeks post-exposure ofAe. aegypti
exposed as first instars to four dosages of SjHe.
Mean Percent Infection
11 + 16(21)
14 + 13 (36)
12 7 (37)
46 41 (24)
58 + 59 (13)
58 + 30 (36)
17 24 (6)
0 + 0 (4)
0 0 (33)
4 6 (24)
33 42 (27)
a Three replicates were conduced at 1 and 3 weeks, one replicate at 2 weeks, total number
of individuals examined across all replicates appears in parentheses.
b Not present.
Table 4-9. Logistic regression of percent infection at 1, 2, and 3 weeks post-exposure.
Variable df X2 P
Week (1 vs 2) 1 0.00 0.9998
Week(l vs 3) 1 1.31 0.2519
Week (2 vs 3) 1 0.00 0.9998
5 x 102
1 x 103
5 x 103
1 x 104
90 C-- control
90 A 1000
8 -- 10000
*# 50 _____ r_ _
0 1 3 5 7 9 11 13 15 17 19 21
Time Post-Exposure (days)
Figure 4-3. Percent mortality over time of first instar Ae. aegypti exposed to two dosages
of SjHe. Two replicates were conducted.
. . .
. .. .
. . . .
. .. . .
7 9 11 13 15 17 19
Time Post-Exposure (Days)
Figure 4-4. Average control proportion of larvae, pupae, and adults of surviving
Ae. aegypti 1-3 weeks post-exposure.
'60 o Adults
.E 50 a Pupae
._ 40 Larvae
7 9 11 13 15 17 19
Time Post-Exposure (Days)
Figure 4-5. Average proportion of larvae, pupae, and adults of surviving Ae. aegypti at 1-
3 weeks post exposure to 1 x 104 cysts/mL of SjHe.
Table 4-10. Mean SD adult male:female ratio ofAe. aegypti at 3 weeks post-exposure
to 2 dosages of SjHe.
Dosage (cysts/mL) Mean M:F Ratio"
0 2.1 0.9a
1 x 103 2.3 + 1.5a
1 x 104 0.8 0.2a
a Means followed by different letters were significantly different (proc glm, Ismeans; P <
Table 4-11. Corrected percent mortality over time ofAe. aegypti at three dosages of SjHe
and four food levels
Days post exposure
Dosage Food 1 2 3 4 5 6 7
1 x 103 0.05 4 0.2a 22 6 35 25 43 18 51 27 53 30 54 36
0.1 3 3 12 17 14 20 14 22 22 31 26 37 30 30
0.2 3+2 3+2 3+5 6+5 11+10 9+9 21+22
0.4 2+3 2+3 6+9 6+9 6+9 6+9 6+9
1 x 104 0.05 12 5 96 6 100 + 0 100 + 0 100 + 0 100 + 0 100 + 0
0.1 22+6 81+10 87+5 92+0 93+2 96+4 98+2
0.2 7+6 53+9 67+11 84 3 84 3 92 7 95 +6
0.4 10+14 23+3 40+9 54 24 57 32 62+30 72 23
1 x 105 0.05 37 27 100 +0 100 +0 100 +0 100 +0 100 +0 100 0
0.1 35+21 100+0 100+0 100+0 100+0 100+0 100+0
0.2 20 1 96 6 98 3 100 +0 100 +0 100 +0 100 0
0.4 16 6 92 +0 94 +3 94 3 95 5 95 6 97 + 5
a Three replicates were performed, Abbott's correction for control mortality was used
d d d dd
6b 0 1.00E+03
50 El 1.00E+04
40 0 1.00E+05
E ab a
0.05 0.1 0.2 0.4
Figure 4-6. Percent mortality at 2 days post-exposure of first instar Ae. aegypti exposed to
three dosages of Helicosporidia and four food levels. Three replicates were
performed. Means followed by different letters were statistically different
(proc glm, Ismeans; P < 0.05). Abbott's correction for control mortality was
Table 4-12. Mean ( SD) corrected percent mortality in Aedes aegypti 7 days after
exposure to SjHe at four different food levels and three dosages of
7-day Percent Mortality
Dosage (cysts/mL) 0.05 0.1 0.2 0.4
1 x 103 43 36a ab 27 25ab 22 19ab 3 6b
1 x 104 93 14a 91 + 16a 77 39a 54 40a
1 x 105 100 + Oa 100 + Oa 99 2a 86 23a
a Three replicates were performed, consisting of 25 larvae each, Abbott's correction for
control mortality was used.
b Means followed by different letters are statistically different within each dose
(proc glm, Ismeans; P < 0.05).
Table 4-13. Mean ( SD) infection rates in surviving Ae. aegypti 7 days after exposure to
three dosages of SjHe at four different food levels. Three replicates were
performed. The total number of individuals examined across all replicates
appears in parentheses. Means followed by different letters were statistically
different (proc glm, Ismeans; P < 0.05).
Dosage (cysts/mL) 0.05 0.1 0.2 0.4
1 x 103 72 25a(24) ab 52 25ab (34) 34+8bc(41) 14 10c(36)
1 x 104 100 +0(2) 83 24 (5) 67 37 (18)
1 105 100 (1) 0 (2)
a Three replicates were performed, consisting of 25 larvae each, Abbott's correction for
control mortality was used.
b Means followed by different letters are statistically different within each dose
(proc glm, Ismeans; P < 0.05).
Table 4-14. Mean (+ SD) fixed, FITC-labeled cysts ingested per Ae. aegypti larva
exposed to 1 x 105 cysts/mL at three food levels. Means followed by different
letters were statistically different (proc glm, Ismeans; P < 0.05).
Food (mg/mL) Cysts per insect
0.1 4.2 4.4 x 103a
0.2 3.1 3.0 x 103ab
0.4 2.0 + 2.0 x 103b
Highly variable bioassay data seem to be the norm with many Helicosporidium
isolates. Fukuda et al. (1976) found that infection rates of An. quadrimaculatus exposed
to a mosquito isolate varied between 17 and 93% among four replicates. Blaske and
Boucias (2004) found that oral challenge of three noctuid species with an isolate of
Helicosporidium from an aquatic weevil produced either no infection or only 50%
infection. However, injection of the noctuids with Helicosporidium consistently resulted
in 100% infection. Initial infection events thus may be the source of the variability in
bioassay data. In our study, cyst viability has also been implicated to play a role in the
variable bioassay results produced with different cyst preparations.
Previous studies examining cyst viability have used a standardized mosquito
bioassay. Hembree (1981), for example, exposed groups of 100 first instar 24-hour old
Ae. aegypti to a range of dosages of Helicosporidium after storage or treatment with
various environmental factors, measuring infection rate as the response variable. Avery
and Undeen (1987b), on the other hand, used groups of 150 first instar An.
quadrimaculatus exposed to a range of dosages of Helicosporidium for 24 hours,
measuring 72-hour mortality as the response variable. In vitro methods for assessing cyst
viability have not been tested before. Since dehiscence is necessary to initiate infection
and directly related to viability, dehiscence rate is a logical choice for assessing cyst
viability. Variation in dehiscence rate of SjHe between cyst preparations and decline of
dehiscence rates during storage may account for the high variability in the bioassay data.
However, the relationship between dehiscence and mortality and infection was not very
strong, indicating that in vitro dehiscence may not be the best indicator of cyst viability.
Also, there may be factors other than cyst viability influencing variation in the bioassay
Helicosporidium has a wide host range. Natural infections have occurred in mites,
cladocerans, Coleoptera, Collembola, Diptera, Lepidoptera, Oligochaeta, and parasitic
trematodes. Helicosporidium is not highly host-specific. For example, Kellen and
Lindegren (1973) transmitted a Helicosporidium isolate obtained from Carpophilus
mutilatus Erichson to 9 other species of Coleoptera, 6 species of Lepidoptera, 3 species of
mites, and 1 species of Diptera. There are only five mosquito host records for
Helicosporidium, but Helicosporidium from mosquitoes has been transmitted to 25 other
species of mosquitoes, three species of Coleoptera, and three species of Lepidoptera.
Previous SjHe host range results indicated that Ae. aegypti was a less suitable host
than An. quadrimaculatus (Conklin et al. 2005). For example, in previous studies, SjHe
infection rates were below 20% for first instar Ae. aegypti treated with 1 x 105 cysts/mL.
In our study, the infection rate of first instar Ae. aegypti exposed to 1 x 105 cysts/mL was
75 25%. Also, in previous studies, mortality of first instar Ae. aegypti exposed to
1 x 105 cysts/mL SjHe was less than 10%. In our study, mortality of first instarAe.
aegypti exposed to 1 x 105 cysts/mL was 83 24%. In addition, Conklin et al. (2005)
reported that the SjHe isolate did not cause melanization in mosquito hosts, whereas the
isolate from an aquatic weevil (CsHe) did cause melanization. In our study, melanization
was more prevalent in Ae. aegypti than An. quadrimaculatus, possibly indicating that Ae.
aegypti are still less suitable hosts for Helicosporidium than An. quadrimaculatus. It is
difficult to say what might have caused these differences. Isolated from a blackfly in
2003, SjHe has been continuously amplified in lepidopteran hosts. The Helicosporidial
cells used by Conklin et al. (2005) had been amplified in lepidopterans for less than a
year, but the Helicosporidial cells used in our study had been in culture in H. zea for 1-2
years. Continued selection by amplification in a heterologous host may have caused a
phenotypic change in the isolate over time. Avery and Undeen (1987b) described changes
in cyst diameter of a pond-water isolate after one passage through a lepidopteran host.
Hembree (1981), likewise, amplified a mosquito isolate in S. exigua and reported a loss
of infectivity after one passage through this heterologous host. Rather than exhibiting a
decrease in infectivity, the SjHe isolate appears to have increased infectivity toward
Ae. aegypti. There is also a possibility that the SjHe isolate was contaminated with the
weevil isolate (CsHe) used by Conklin et al. (2005).
Age-dependent resistance to pathogens is common in insects as well as many other
organisms. There are several possible reasons for decreased susceptibility due to age. Age
may alter ingestion or feeding as the larvae become satiated and cease feeding or begin
molting into the next instar. In this study, 24-hour-old first instar larvae molted into the
second instar during the exposure period, possibly resulting in reduced ingestion. Older
larvae also have had time to establish a compliment of immune peptides and phagocytic
hemocytes that may kill invasive microorganisms. In mosquito larvae, the peritrophic
matrix may also be a key to pathogen resistance. Mosquito larvae have a highly
structured peritrophic matrix secreted by a ring of cells in the cardia and extending
through the entire midgut. The peritrophic matrix is already present in neonate
Ae. aegypti, but it is thinner, without the strengthening fibers present in the peritrophic
matrix of later instars. Thickening of the peritrophic matrix occurs as the larvae age and
feed (Richards and Richards 1971).
Decreasing susceptibility with age has been reported for three other
Helicosporidium isolates. Fukuda et al. (1976) reported a 3-fold reduction in infection
rate from larvae 1 to 3 days old. Hembree (1981) found that both instar and age during
the first instar had an effect on infectivity and mortality. In Hembree's (1981) study, there
was a 3-fold difference in IC5o between first and second instars ofAe. aegypti. Seif and
Rifaat (2001) reported that the IC50 of second instar Cx. pipiens was only 1.2 times more
than the first instars. In our study, there was an 8 to 13-fold difference in infection rate
between the first and second instars ofAe. aegypti. The results for the effect of age within
the first instar in this study were similar to those of Hembree (1981), who found a 2-fold
difference in infection rate of 2 and 24-hour old Ae. aegypti at 5 x 102 cysts/mL.
However, the decrease in susceptibility between 2 and 24-hour old Ae. aegypti in our
study was more pronounced, decreasing from 40 to 0% from 2 to 24-hours of age at a
dosage of 5 x 103 cysts/mL. Increasing the dosage of SjHe minimized the effect of first
instar age on susceptibility, indicating that the decrease in susceptibility due to age was
Infection with Helicosporidium reaches its peak just after 7 days post-exposure.
Fukuda et al. (1976) reported that infection could be detected at 4 days post-exposure and
that the maximum number of infected mosquitoes was collected 9-10 days post-exposure.
Hembree (1981) found that virtually all infections could be detected 4 days after
infection, and that 10-14 day-old larvae were optimum for cyst purification. Seif and
Rifaat (2001) reported 5 days post-exposure for the first detectable infections, and that
the maximum number of infected larvae was collected 8-10 days post-exposure. All of
the above authors noted that mortality occurred in the late fourth instar, pupae, and
adults. The pathological effects of Helicosporidium infection manifest themselves at the
larval-pupal interface in lepidopteran hosts also. Blaske and Boucias (2004) found that
infected noctuids successfully pupated, but most died as pupae. A small number of
infected larvae emerged as infected adults. Infected noctuiid adults and pupae were often
malformed. In bioassays with SjHe, percent infection did not change significantly after
1 week post-exposure, and most infected insects died late in the fourth instar, at the
larval-pupal interface. No malformation of pupae or adults was observed in SjHe-treated
Ae. aegypti. Sex-ratio in infected noctuiids was not different from controls (Blaske and
Boucias 2004). Sex ratio of exposed Ae. aegypti also did not differ from controls. The
minor female-bias in the sex ratio ofAe. aegypti exposed to SjHe is due to the high
mortality in these groups.
Infected larvae that successfully pupated and emerged as adults were rare in this
study. Horizontal transmission appears to be the most likely form of transmission for
Helicosporidium. Blaske and Boucias (2004) found that Helicosporidium infection could
be vertically transmitted by infected noctuiid adults. However, the infection rate in the
offspring was very low, and the ovarian tissue itself was not found to be infected. The
role of vertical transmission in mosquitoes is unknown. The only study of vertical
transmission in mosquitoes was by Fukuda et al. (1976), which examined 2060 larvae
from 17 egg rafts of infected Cx. salinarius, but found no infection. Natural horizontal
transmission of Helicosporidium from infected individuals to uninfected individuals of
the same species has not yet been documented with any host species. However, Hembree
(1981) infected Toxorhynchites splendens (Wied.) by feeding the predatory larvae
Helicosporidium-infected Cx. pipiens. Once cysts are produced in the host, they must be
released into the environment in order to be ingested by the next host, but no mechanism
of cyst release has been found (Blaske-Lietze et al. in press).
While death of infected larvae during the fourth instar appears to be common for
Helicosporidium infections, early mortality accounted for most of the total mortality at
7 days post exposure. High early mortality due to exposure to SjHe in this study and also
observed by Conklin et al. (2005) has only been reported for one other isolate of
Helicosporidium. Avery and Undeen (1987b) found that a pond water isolate of
Helicosporidium caused high mortality in first instar larvae at 72-hours post exposure.
The pond water isolate also caused melanization in mosquitoes. Since early mortality
causes the death of the host before cyst production, terminating the infection cycle, the
authors suggested that mosquitoes were not natural hosts of the pond water isolate.
Increasing the food during exposure to SjHe decreased the amount of cysts
ingested, as indicated with fixed, FITC-labeled cysts. However, decreased ingestion may
not completely explain decreased infection. In addition to reduced ingestion, plant
chemicals present in the mosquito diet may have damaged the ingested helicosporidial
cells. Alfalfa, a component of the larval food, produces isolfavonoid phytoalexins,
defensive compounds that exhibit antimicrobial activity in vitro (He and Dixon 2000).
Differences in food composition may account for variation in bioassay results from
previous studies. For example, Hembree (1981) used a high-protein mixture of yeast,
liver powder, and hog chow as larval food, while Fukuda et al. (1976) used rabbit chow,
and Seif and Rifaat (2001) used powdered tropical fish food.
In our study, there was no effect of infection on development. However, there are
other reports that Helicosporidium can influence development in other hosts. For
example, Conklin et al. (2005) found that the mean head capsule measurements of
Ae. aegypti exposed to SjHe and reared in groups were smaller than the control larvae.
However, this approach failed to take into account the effect of group size on host
development. Individual rearing eliminates this larval density factor, providing each larva
with nearly ideal conditions for growth. It is possible that, in groups, larval density and
competition between larvae for food increases the cost of infection, thus increasing the
effect of infection on development. In support of this, Bedhomme et al. (1999) found that
Ae. aegypti infected with a microsporidian developed more slowly when placed in groups
than individually-reared larvae.
Utilization of the time to pupation as a measurement of development may not have
been adequate. Blaske and Boucias (2004) reported that the effect of infection on noctuid
development varied by species and measurement of development. Pupal weight, time to
pupation, and time to eclosion each provided a different profile of the effect of
Helicosporidium infection on development. Also, analysis of time to pupation in SjHe-
infected Ae. aegypti would have been improved by recording pupation more than once
every 24 hours. With such a short larval period, the difference in development between
infected and control mosquitoes may have been less than 1 day. However, Hembree
(1981) reported that pupation was delayed 1-2 days in infected Ae. aegypti and pupation
was lengthened. Different rearing conditions may have caused slower development in his
infected larvae. The length of time to development (pupation or adulthood) can also have
an effect on a pathogen's ability to influence host development. The length of the larval
period of Ae. aegypti at 26 C is 7-10 days, while the length of the larval period of the
weevil D. abbreviatus is 100 days or more (Conklin et al. 2005), providing the pathogen
a greater opportunity to influence the development of the host. Accordingly, there was a
pronounced effect on host development in D. abbreviatus treated with SjHe.
While Ae. aegypti and An. quadrimaculatus support vegetative growth of SjHe,
several aspects of the interaction between SjHe and mosquitoes indicate that host
suitability is low for these species. The IC5o for first-instar An. quadrimaculatus was
between 1 x 104 and 1 x 105 cysts/mL (Conklin et al. 2005), and forAe. aegypti the IC50
of first-instars was also in the range of 1 x 104 cysts/mL. The dose-response of two
culicid isolates have been examined using similar methods, with much lower ICso's than
this study. Hembree (1981) found that the IC5o of first instarAe. aegypti exposed to a
mosquito isolate from Thailand was approximately 1 x 103 cysts/mL. Seif and Rifaat
(2001) found that for first instar Cx. pipiens exposed to another mosquito isolate, the IC5o
was 1.9 x 102 cysts/mL. High initial mortality ofAe. aegypti and An. quadrimaculatus
exposed to SjHe limited the number of infected, cyst-producing larvae recovered later.
Cyst production after death also does not appear to occur in other hosts of
Helicosporidium. Blaske-Lietze et al. (in press) have found that in H. zea, vegetative
reproduction halts after the host's death, and further cyst differentiation does not occur in
the dead host. This interaction between a pathogen and the non-host species did not
benefit either the host or the pathogen. The host had an acute reaction, causing death and
shutting down pathogen reproduction, while the pathogen, having prematurely killed its
host, was unable to complete its life cycle.
The host range of a single Helicosporidium isolate may be deceptively broad,
where many species can be infected, but only a few support a high level of replication
and transmission, representing true host species. The sensitivity of the bioassay system of
SjHe and mosquitoes to cyst viability, minor changes in host age, and food availability
also indicate a poorly adapted host-pathogen interaction.
Table A-1. Compiled cyst measurements.
References Size (unm) Stain Fixation Host
Keilin 1921 5 6 None None Dasyhelea obscura
Iron- Camoy's fluid,
Weiser 1970 3 4.5 haematoxylin paraffin Hepialis pallens
Iron- fluid + 1%
Weiser 1970 4.5 5 haematoxylin HAc Dasyhelea obscura
Hembree 1979 5.5 Giemsa Methanol Ae. u, --' pti
Hembree 1979 5.9 None None Ae. u, --'/i
Kellen & Lindegren Paramyelois
1974 5.03 + 0.28 Giemsa Methanol transitella
Kellen & Lindegren Paramyelois
1974 5.59 + 0.26 None None transitella
Sayre & Clark 1978 5.6 + 0.052 Giemsa None Daphnia magna
Pekkarinen 1993 5.31 + 0.04 None Glutaraldehyde Bucephalid trematode
Pekkarinen 1993 4.99 + 0.04 Stained None Bucephalid trematode
Pekkarinen 1993 4.91 + 0.05 Stained None Bucephalid trematode
Pekkarinen 1993 4.92 + 0.06 Stained None Bucephalid trematode
Pekkarinen 1993 3.71 -5.14 TEM TEM Bucephalid trematode
6.5 + 0.2 x
Boucias et al. 2001 5.9 0.3 None None Simulium jonesi
6.2 + 0.3 x
Boucias et al. 2001 5.9 + 0.1 None None Helicoverpa zea
Avery & Undeen
1987a 7.19 + 0.15 Giemsa None Collembolan
Avery & Undeen
1987a 5.58 + 0.03 None None Helicoverpa zea
Avery & Undeen
1987b 8.91 + 0.12 None None Collembolan
Table A-2. Compiled filamentous cell measurements
Authors Size ([tm) Stain Mount Host
Keilin 1921 60 65 x 1 None None Dasyhelea obscure
Kellen & Lindegren
1974 50.4 + 3.32 None Saline Paramyelois transitella
Sayre & Clark 1978 61.7 2.44 Giemsa None Daphnia magna
37 4.3 x
Boucias et al. 2001 0.9 + 0.13 None None Simulium jonesi
CONSTRUCTION OF MOSQUITO BREEDERS
When rearing mosquitoes to adulthood, it becomes necessary to find a way to have
both water for the larvae and pupae and space for the adults to be collected after
emergence. Placing the larval rearing pan into a large screen cage is one simple way to
address this problem. However, screen cages are bulky and difficult to sterilize, and
collecting adults from such a large space can be a challenge. Many "mosquito breeder"
products that address this problem are available through mail order, but these usually cost
$10-15 each. A mosquito breeder has a relatively simple design, however, and can be
constructed from materials that most laboratories have on hand. In this series of
experiments, mosquito breeders were constructed from 16 oz clear, plastic food
containers with tight-fitting lids ("Del-Pak" by Reynolds Grant Park, IL). These
containers are light, durable, dishwasher-safe, stackable, water-tight, easy to cut with a
razorblade, and recyclable. The following is a protocol for constructing mosquito
* Three 16 oz plastic tubs
* Two lids for tubs
* One 5x5 in piece of green tulle
* Hot glue gun and glue sticks
1. Using the razorblade, make an X in the middle of one of the lids. Pop this out and
fold back the flaps. Cut one of the flaps off to facilitate movement of adults out of
2. Cut out the middle of the other lid, leaving only the lip that attaches to the
container. Do this by making a slit with the razorblade, then finishing cutting with
3. Heat up the hot glue gun and glue the two lids together top to top. Run a bead of
glue all the way around the outside edges.
4. Cut the bottoms off of two of the plastic tubs using the razorblade.
5. Place one of the bottomless tubs upside down and place the tulle on top of the
6. Press the other bottomless tub upside down on top of the other tub, securing the
tulle between the two tubs.
7. Assemble the finished mosquito breeder by snapping the remaining tub and the
tulle part into the two sides of the lid (Figure B-l).
Each finished mosquito breeder can hold about 500 mL of water in the bottom
container. A cotton ball soaked in sugar solution can be placed on top of the tulle screen
to provide food for the adults. Collecting adults from the container is accomplished by
placing a lid with a hole punched in the middle over the top of the screening. Introduce a
tube through the hole and pump CO2 into the container for approximately 1 min, or until
all mosquitoes are knocked down. The mosquitoes can be collected with forceps. Not all
adult mosquitoes will fly up into the upper part of the breeder, so the lid will have to be
removed to collect those that remain below. The approximate cost of each mosquito
breeder constructed as above is 41 cents.
Figure B-1. Finished mosquito breeder
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Tracy Conklin was born on April 17, 1982 in West Palm Beach, Florida. As an
only child, she entertained herself by catching insects, lizards, and fish in her subtropical
backyard. She examined her first mosquito larva under her mother's microscope at 8
years old. Little did she know that she would spend 3 years of her life doing the same in
college. After spending 2 years of her undergraduate career studying British literature,
Tracy gave in to her love for insects and changed her major to entomology. As an
undergraduate, she worked for Dr. Drion Boucias in the insect pathology laboratory, and
after graduating, decided to continue looking at mosquitoes under the microscope.