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Comparative development and tissue tropism of Nosema apis and Nosema ceranae
Accepted Manuscript Comparative development and tissue tropism in Nosema apis and Nosema ce‐ ranae Wei-Fone Huang, Leellen F. Solter PII: DOI: Reference:
S0022-2011(13)00004-9 http://dx.doi.org/10.1016/j.jip.2013.01.001 YJIPA 6401
To appear in:
Journal of Invertebrate Pathology
Received Date: Accepted Date:
16 October 2012 4 January 2013
Please cite this article as: Huang, W-F., Solter, L.F., Comparative development and tissue tropism in Nosema apis and Nosema ceranae, Journal of Invertebrate Pathology (2013), doi: http://dx.doi.org/10.1016/j.jip.2013.01.001
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Comparative development and tissue tropism in Nosema apis and Nosema ceranae
2 3 4 5
Wei-Fone Huang* and Leellen F. Solter
6 7
Illinois Natural History Survey, Prairie Research Institute, University of Illinois
8
1816 S. Oak Street, Champaign, IL 61801 USA
9 10 11 12 13 14 15
*Corresponding author: [email protected]
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Abstract
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The two etiological agents of nosema disease in honey bees, Nosema apis and Nosema ceranae
19
(Microsporidia: Nosematidae), reproduce in the midgut tissues of the host. Nosema apis is tissue
20
specific but the development and tissue tropism of Nosema ceranae is not well understood. Our
21
investigations compared development of the two phylogenetically related pathogens in all major
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host tissues. Using microscopy, PCR and qPCR quantification to evaluate tissue tropism of
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infected bees in communal cages and of individually restrained infected bees, we found no
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detectable spores in cephalic or other body tissues except midgut tissues. Nosema DNA was
25
detected in Malpighian tubules but the tubules could not be separated from the alimentary tract
26
without release of spores from the midgut. Nosema DNA was not detected in hemolymph
27
sampled from the head capsule or the abdomen of infected bees. We confirmed that N. ceranae
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only develops in midgut tissues. Spores of both species released from host midgut cells
29
accumulated in the hindgut lumen, and we noted differences in numbers and ratios of spore types
30
and in growth curves between the two pathogens. Nosema apis reached a consistent level of
31
spore production after 12 days post inoculation (dpi); Nosema ceranae spore production
32
increased linearly from 12-20 dpi and the number of mature Nosema ceranae spores was
33
consistently higher.
34 35 36
Keywords: Nosema; honey bee; tissue tropism; glands; transmission; development
37 38 39 40
Introduction
41
Apis mellifera, and cause chronic, debilitating disease. Infections frequently occur in high
42
prevalence in apiaries and deleteriously impact pollination services and production of honey
43
(Bailey and Ball, 1991; Fries, 1993). N. apis was the only known agent of honey bee nosemosis
44
until N. ceranae, a genetically related but distinct species, was described by Fries (1996) from
45
the Asiatic honey bee Apis cerana. N. ceranae was first reported infecting A. mellifera in Taiwan
46
in 2005 (Huang et al., 2007) and then in Europe (Higes et al., 2006), and was subsequently
47
identified in A. mellifera globally (Huang et al., 2008; Klee et al., 2007). The timing and means
48
of the invasion events are not known; N. ceranae was the predominant microsporidian pathogen
49
in A. mellifera by the mid-1990s in North America (Chen and Huang, 2010) and before 1990 in
50
Uruguay (Invernizzi et al. 2009).
51
Two microsporidian pathogens, Nosema apis and Nosema ceranae, infect the western honey bee,
Early morphological descriptions of N. apis were vague (McIvor and Malone, 1995), but
52
histological observations and tissue tropism studies were consistent: N. apis only infects the adult
53
stage of the host, and only reproduces in the midgut epithelium cells (Bailey and Ball, 1991; De
54
Graaf and Jacobs, 1991; Fries, 1993). The strict tissue tropism of N. apis is uncommon in the
55
genus. Most related Nosema species, including the closely related Nosema bombi infecting
56
bumble bees (McIvor and Malone, 1995; Fries et al., 2001), are systemic pathogens that
57
reproduce in the cells of most body tissues, including the gonads, and many species are
58
transmitted transovarially from infected females to their offspring (Becnel and Andreadis, 1999).
59
N. apis infection in queens may affect the development of oocytes (Liu, 1992) but the pathogen
60
is not transovarially transmitted (Webster et al. 2008).
61
N. ceranae, like N. apis, was described as a midgut pathogen of adult bees (Fries, 1996;
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Fries 2010) and the two species are morphologically quite similar (Fries et al., 1996); therefore,
63
PCR has become the primary tool for differential diagnosis of honey bee nosematosis. When
64
PCR methods were used to detect N. ceranae, DNA signals were also observed in the cephalic
65
tissues (Chen et al., 2009; Copley and Jabaji, 2012; Gisder et al., 2010), but neither vegetative
66
forms nor mature spores have been observed in cephalic tissue smears and stains (Gisder et al.,
67
2010). Transmission electron microscopy showed changes in hypopharyngeal gland development
68
and cell micro-morphology in N. apis-inoculated bees but, like N. ceranae, no observable
69
infection (Liu, 1990). The possibility that N. ceranae infects hypopharyngeal and salivary glands
70
potentially changes the interpretation of transmission routes, pathogenesis, and host behavior,
71
and a reservoir function was suggested for the cephalic glands of both Nosema species (Copley
72
and Jabaji, 2012).
73
N. ceranae appears to have replaced or at least successfully competed with N. apis in
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many, if not most, A. mellifera populations globally (Chen et al., 2009; Chen et al., 2012;
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Martin-Hernandez, 2012). To address the characteristics of N. ceranae that have enabled it to
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out-compete N. apis, an established pathogen, we compared the rate of development in the host,
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production of spores, and tissue tropism. We used PCR and qPCR diagnoses for detection of N.
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ceranae and N. apis in the tissues but rearing methods and dissection techniques were newly
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designed to reduce the possibility that potentially uninfected target tissues were contaminated
80
with spores or DNA.
81 82
Materials and Methods
83
Nosema spp. isolates
84
Nosema ceranae was originally collected from campus apiaries at the University of Illinois at
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Urbana-Champaign, and N. apis spores were provided by T. Webster at Kentucky State
86
University. To produce infective spores for experiments, honey bees were held in plastic cages
87
consisting of 480 ml HDPE lidded plastic cups with 3-mm hardware cloth tops (Webster et al.,
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2004) and were inoculated with 105 spores of N. apis or N. ceranae. Bees were anesthetized on
89
ice, then fastened, dorsal side down, on foam boards with insect pins (Fig. 1). Upon warming,
90
bees were individually inoculated by allowing them to imbibe 2 µl sugar water containing 105 N.
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apis or N. ceranae spores, a dosage that produces 100% infection (unpublished data). Bees were
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returned to cages and at ≥10 days post inoculation (dpi), midgut tissues of infected bees were
93
excised, homogenized in a glass tissue grinder, and spores were isolated in sterile water
94
suspensions by centrifugation (Huang et al., 2007). Number and maturity of isolated spores were
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checked under phase-contrast microscopy (400 X) using a Petroff-Hausser counting chamber.
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Only freshly isolated spores were used in bioassays.
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Inoculation of experimental bees
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Newly emerged bees were harvested daily from a brood frame selected from a fumagillin-free
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hive and were held in growth chambers at 34°C for 5 days. The bees were fed 50% sugar water
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and pollen patties (MegaBee™, 15% pollen) ad libitum. Experimental bees were starved for 2 hr
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and inoculated as described above. Only bees ingesting the full 2 µl suspension were included in
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the trials. After allowing the bees to rest on the foam board for 30 min, they were transferred into
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cages as described above, 30 bees per cage, and held in a 30°C growth chamber, 65% relative
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humidity, 24 hr dark. The rearing temperature was reduced to 30°C for experiments because our
105
studies have shown that more intense infections develop at this temperature and high
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temperatures may have adverse effects on infections (Martin-Hernandez et al., 2009; Chen et al.,
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2012). In addition, our results are comparable to previous studies (Forsgren and Fries, 2010;
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Paxton et al., 2007).
109
Examination of tissues
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Honey bees were held for 14-15 days in communal cages as described above. Treated bees were
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rinsed in 0.09% sodium hypochlorite (3% household bleach) with 0.2% tween20, then two rinses
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in sterile water, and individually dissected. Tissues, including the midgut (with intense infection
113
levels), Malpighian tubules, fat body, crop and cephalic tissues, were excised and rinsed in PBS.
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Small tissue samples were smeared on slides and examined under 400x phase contrast
115
microscopy. Giemsa stains were made of fresh tissue samples (Becnel, 2012) and were examined
116
under 400x bright field microscopy.
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Spore counts
118
We observed spores in the hindgut contents of infected bees under phase-contrast microscopy
119
with refringency that was intermediate between the bright refractivity of mature “environmental”
120
(infective) spores and empty (germinated) spores (Solter et al., 2012). Based the morphology of
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spores at specific time periods (Solter and Maddox, 1998; Higes, 2007; Solter et al., 2012) we
122
assumed that intermediately refringent spores with visible polar vacuoles are internally infective
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“primary” spores (Fries et al., 1992; Solter et al. 2012). Other developing spores, e.g. immature
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environmental spores, have similar refringency, although the polar vacuole is usually not visible.
125
Because it is even more difficult to distinguish spore stages and maturity based on refringency in
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cell counting chambers (Petroff-Hausser), spores with similar refringency were assigned to the
127
same category. We thus counted three categories of spores, mature spores (high refringency),
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primary/immature spores (low to medium refringency), and germinated spores (spore husks with
129
little refringency), every 2 days over the course of the test period. We conducted three trials; two
130
trials also served as inoculated positive control sets for our previous research (Huang et al.,
131
submitted).
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Restraint of individual honey bees
133
We designed a restraint system to isolate test bees from colony members and eliminate grooming
134
and other hygienic behaviors. The tapered ends of 500-µl plastic microcentrifuge tubes were
135
removed, leaving an opening in the bottom of the tubes slightly larger than the diameter of the
136
head capsule but smaller than the thorax of A. mellifera. An anesthetized bee was placed head-
137
first into each tube, ensuring that the head emerged through the opening. Cotton was placed in
138
the remaining space to restrain the bee and the top was closed (Fig. 2A). After the bees warmed
139
and became active, they were inoculated with 105 N. ceranae spores in 2 µl water with a
140
micropipetter. After 30 min, the bees were transferred in the restraints to a 1.5 ml
141
microcentrifuge tube with 50% sugar water in the bottom that the bee could reach with its tongue
142
(Fig. 2B). The feeder was replaced every two days. The restrained bees were held in growth
143
chambers under the conditions described above and were dissected at 14 dpi to evaluate
144
infections in the midgut and cephalic tissues. We used either conventional dissection methods or
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dissection through the head capsule vertex (see below) to dissect the restrained bees. The sugar
146
water removed from feeders 12 and 14 dpi was collected and diluted by adding ddH2O to remove
147
the sugars, to a total of 500 µl. The suspension was centrifuged and the supernatant was
148
discarded by pipetting, leaving approximately 10 µl of suspension. The dilution procedure was
149
repeated and the remaining 10-µl suspension was vortexed and observed under 400x phase
150
contrast microscopy, and DNA was extracted for PCR according to the following method.
151
Tissue sampling for PCR
152
Prior to surface sterilization of head capsules, the mouthparts of N. ceranae and N. apis
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inoculated bees were individually sampled by pipetting 5 µl PBS several times between the
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mandibles. DNA was then extracted from the rinsate. Both the communally caged and
155
individually restrained bees were sampled.
156
Bees were individually surface sterilized by washing them twice in cold 0.09% sodium
157
hypochorite with 0.2% tween20, and then rinsing twice in sterile water with 0.2% tween20. Bees
158
were dried on a clean paper towel and held on ice prior to dissection. Cephalic tissues, brain
159
(nerve tissues), hypopharyngeal glands and salivary glands of bees infected with either Nosema
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species were collected, as well as foregut, midgut, Malpighian tubules, second tergum (with
161
attached tissues), and venom gland, of N. ceranae inoculated bees. Tissue smears were made
162
from small samples of fresh tissue and examined under 400x phase contrast microscopy. At least
163
one of the paired cephalic glands and approximately 2 mm3 cubic tissues of brain of each bee
164
were used for the DNA extractions.
165
The conventional method for excision of cephalic tissues we used was similar to that used
166
in published studies (Chen and Huang, 2010), and is briefly described as follows. The head was
167
removed with clean scissors and transferred to a Petri dish containing a few drops of PBS. The
168
cuticle was carefully cut through the oral cavity without touching internal tissues.
169
Hypopharyngeal glands, salivary glands and brain tissues were removed with clean forceps and
170
the tissues were washed twice in PBS. The tissues were transferred to a PCR tube containing
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DNA extraction buffer. In addition to the tissues, 5 µl PBS (with no observable tissues) were
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removed from the Petri dish and were added to DNA extraction buffer to determine if spores
173
were shed into the buffer. DNA was extracted for qPCR.
174
To prevent possible contamination with Nosema sp. spores during dissection, a second
175
method was used to excise cephalic tissues. Anesthetized, surface sterilized bees were dissected
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without decapitation; the head was opened directly through the vertex area without touching the
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mouthparts and pharynx. The cuticle was carefully removed, cutting along the vertical sutures
178
and sutures around the compound eyes, then tissues were removed from directly from the
179
opening with fine forceps. The tissues were placed in a few drops of PBS in a Petri dish and
180
separated under stereomicroscopy, then washed in PBS and transferred to DNA extraction buffer.
181
DNA was extracted as previously described. The PBS rinse was also sampled to check for shed
182
spores as previously described.
183
We sampled hemolymph from the head and abdomen of several infected surface
184
sterilized bees during dissection of tissues. Approximately 2 µl hemolymph was obtained from
185
the head capsule when dissecting via the vertex, and the same amount of abdominal hemolymph
186
was obtained via a razor cut of intersegmental membrane between the first and second abdominal
187
segments. DNA was extracted from hemolymph samples as described below.
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DNA extraction
189
DNA was extracted using a modified Chelex method (Cordes et al., 2012). Dissected tissues,
190
approximately 2 mm3 per sample, were transferred into 50 µl Chelex buffer, 5% Chelex 100
191
(Biorad), 1ng/ul proteinase K and 5% tween20 in ddH2O, in a 200 µl PCR tube. The tubes were
192
vortexed at high speed for 15 s and centrifuged briefly to precipitate the Chelex buffer and tissue
193
to the bottom of the vial, then incubated using the following program: 56°C for 2 hr, followed by
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95°C for 30 min, and a 4°C cool-down. The homogenate was centrifuged at 13,000 g for 10 min
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and the supernatant retained for PCR.
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qPCR and PCR
197
Quantitative PCR was performed using the SYBR green method. The specific primer set for N.
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ceranae was Nc841f: GAG AGA ACG GTT TTT TGT TTG AGA; Nc980r: ATC CTT TCC
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TTC CTA CAC TGA TTG, estimated amplicon size 140 bp. For N. apis, primers Na65f: CGT
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ACT ATG TAC TGA AAG ATG GAC TGC and Na181r: AGG TCT CAC TCT TAC TGT
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ACA TAT GTT AGC, estimated amplicon size of 116 bp, were designed using the SSUrRNA
202
sequence alignment. The reactions were carried out on an ABI 7900HT thermal cycler with
203
standard two-step PCR method using SYBR® Green PCR Master Mix (ABI). The annealing
204
temperature was 64.5C with 40 cycles. No cross-reaction between N. apis and N. ceranae or
205
non-specific PCR product was found. The results were analyzed using absolute quantification set
206
up in SDS 2.2 (ABI) with spore extractions as standards.
207
Spore extractions that were filtered, centrifuged, re-suspended, and counted were used as
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standards. Four standard samples of both N. apis and N. ceranae in log dilutions of 5x10 to
209
5x104 spores were used to construct the standard curves for qPCR. We used identical methods
210
and volume for the standards and dissected tissues, and extracted the DNA at the same time. The
211
same primer sets were used for regular PCR diagnosis with the same reaction program, but with
212
Platinum® Taq (Invitrogen) following manufacturer’s directions.
213
Statistical analysis
214
The spore counts were analyzed by one-way ANOVA to determine significant differences using
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SPSS software (IBM).
216 217
Results
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Spores in hindgut lumen contents
219
Spore counts from the hindgut lumen contents are shown in Figure 3. Total numbers of the three
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types of spores, mature, immature/primary, and germinated, were all significantly different
221
(P<0.001) between N. ceranae and N. apis over the course of the infection period. More mature
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spores and fewer germinated and primary/immature spores were observed for N. ceranae than
223
for N. apis. We analyzed the ratio of types of spores in hindgut contents at 12 dpi and found 80-
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95% mature N ceranae spores, while 57-83% of N. apis spores were mature. The ratio of N. apis
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mature spores increased during the infection period (Fig. 3). The growth curve for germinated
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spores was irregular in N. apis infections, but increased linearly to 16 dpi in N. ceranae
227
infections. The total number of primary/immature spores increased to 14 dpi and then dropped
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rapidly for both N. apis and N. ceranae infections.
229 230
Microscopic and PCR evaluation of tissues from caged bees
231
Midgut spore counts exceeded 106 spores in all treated bees at 14 dpi for both N. ceranae and N.
232
apis. No spores or vegetative stages were observed in tissues other than the alimentary tract,
233
however, we observed spores attached to the surfaces of dissected tissues, including Malpighian
234
tubules and the tissues within the tergum. PCR diagnoses produced signals for most tissues but
235
the microsporidia did not appear to be developing within the cytoplasm of host cells. We noted
236
that heavily infected midgut tissues are extremely fragile and break down during dissection, but
237
microsporidia were not observed in the hemolymph extracted prior to the dissection of the hosts.
238
The efficiency of qPCR to quantify spores from DNA was in the range of 90-98% for N.
239
apis and 96-99% for N. ceranae. Six qPCR tests (three repeats for each test, three tests for each
240
sample) on different sample sets were performed to generate the results in Table 1. Standard
241
curve and PCR efficiencies were linear and consistent among the different reactions.
242
Quantitative PCR measurements for Nosema DNA in tissues were inconsistent for
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infected communally caged honey bees that were dissected using the conventional method
244
(Table 1). Three different cephalic tissues, brain, salivary glands and hypopharyngeal glands
245
were compared, and no N. ceranae DNA was detected in brain tissues of six of the infected bees.
246
Because inconsistent DNA quantities were found in cephalic tissues among bees, we could not
247
evaluate differences among these tissues. Tissues from N. apis-inoculated bees produced similar
248
results, but we detected N. apis DNA in brain tissues of two of four inoculated bees. All PBS
249
rinsate remaining from the conventional dissections contained Nosema DNA. The rinsate from
250
external mouthparts, mandibles and partial labium contained several thousand spores based on
251
qPCR results.
252
There was no detectable N. ceranae or N. apis DNA in the cephalic tissues or in the PBS
253
used to rinse and separate tissues dissected through the head capsule vertex. The same group of
254
bees was used for both types of dissection, but an additional N. ceranae inoculation trial using
255
the vertex dissection method confirmed the results. There was no detectable Nosema DNA in the
256
hemolymph samples from caged bees.
257 258
Infections in restrained bees
259
We restrained and isolated bees only for N. ceranae inoculations. Mortality was higher for
260
individually restrained bees during the course of the experiment (75%), than for caged bees
261
(approximately 10%), most likely due to isolation and restriction to movement, and midgut spore
262
counts were lower. Caged bees fed the same inoculum had 5-37 million spores in midgut, while
263
restrained bees produced 1-3 million spores. Twelve restrained honey bees survived to 14 dpi.
264
Infections in caged bees were late stage and well-developed. Infections in restrained bees
265
consisted of predominantly mature environmental spores but the mortality rate suggests that bees
266
with extremely high levels of infection did not survive. No microsporidian stages were observed
267
in cephalic tissues under phase contrast microscopy or in qPCR diagnosis of these tissues that
268
were dissected from restrained bees using the conventional method or the dissection method via
269
the head capsule vertex. We found no spores in mouthpart rinses or sugar water at 12 and 14 dpi.
270 271
Discussion
272
We compared the development of infection based on spore accumulation in the hindgut, and
273
tissue tropism of the two related microsporidian pathogens infecting the western honey bee. In
274
previous studies, no spore types were distinguished and spores were counted in the midgut or in
275
the entire abdomen (midgut tissue and hindgut contents) (Forsgren and Fries, 2010; Martin-
276
Hernandez et al., 2009; Paxton et al., 2007), although both primary spores and empty
277
(germinated) spores can be observed under phase contrast microcopy (Solter et al., 2012). Most
278
bees we used for these experiments avoided defecating inside the cages, so the change in ratio of
279
the different spore types accumulated in the hindgut reflected the progress of infection in the
280
midgut. PCR-based diagnosis cannot distinguish primary, immature and mature environmental
281
spores. We noted a significantly lower ratio of primary/immature spores in the hindgut contents
282
of N. ceranae-infected bees than of N. apis-infected bees. Primary spores and, we assume,
283
immature environmental spores, do not survive outside the host and do not appear to be
284
transmissible among hosts, thus the higher ratio of environmental spores potentially provides N.
285
ceranae a competitive advantage. The lower primary/immature spore ratio in N. ceranae
286
infections could imply better suppression of host cell immune response, resulting in lysis of
287
midgut cells with more advanced infections and more mature spores than occurs in N. apis
288
infections. The ratio of environmental spores was high in both species after 14 dpi.
289
Our observations of mature environmental spores in host hindgut contents suggested that
290
proliferation rates differed for N. ceranae (approximately 8×106 spores/day), and N. apis
291
(approximately 4×106 spores/day) based on estimated linear growth curve slopes after the spore
292
count in the midgut tissues reached the plateau phase at 10-12 dpi (Forsgren and Fries, 2010;
293
Paxton et al., 2007). Spore accumulation in the hindgut trended higher for N. ceranae than N.
294
apis after 12 dpi on a per sample-day basis and was significantly higher overall. N. ceranae
295
spores appear to mature later in the infection process than do N. apis spores. This observation
296
corresponds to TEM observations of midgut infections (Higes et al., 2007).
297
Previous research detected DNA but no observable microsporidian stages in cephalic
298
tissues of N. apis infected bees, therefore we investigated possible cephalic infection by N.
299
ceranae using qPCR to evaluate infection development. We used different dissection and caging
300
methods (communal or isolated) to eliminate the possibility of contamination. DNA of both N.
301
apis and N. ceranae was detected by qPCR in cephalic tissues when conventional dissection
302
methods via the mouthparts were used, but excision of cephalic tissues directly though vertex of
303
the head capsule of both cage-reared and restrained bees eliminated touching of the mouthparts,
304
which could be a source of contamination in qPCR detections. No N. apis or N. ceranae DNA
305
was detected in these qPCR samples. Moreover, we found no detectable N. ceranae DNA in
306
hemolymph collected from the head or abdomen, suggesting the inability of N. ceranae to enter
307
the hemocoel as was also determined for N. apis using an antiserum technique (De Graaf and
308
Jacobs, 1991). Our results do not support the hypothesis that N. apis and N. ceranae infect the
309
honey bee cephalic glands and use them as reservoir tissues (Copley and Jabaji, 2012). Spores
310
and DNA of both N. apis and N. ceranae were found only in the alimentary tract.
311
The mouthparts of infected caged bees contained relatively high numbers of spores,
312
possibly due to hygienic grooming in cages holding multiple infected bees, some of which likely
313
defecated in the cages, releasing spores. Fecal matter not removed from mouthparts by surface
314
sterilization could, thus, be a source of contamination during conventional dissections of cephalic
315
tissues. Although we did not observe fecal material on cage surfaces, we did find spores of both
316
Nosema species in rinsate brushed from the cage walls. No spores were found in the mouthparts
317
or food source of individually restrained infected bees. Restrained bees were prevented from
318
performing hygienic behaviors and we suggest that presence of spores in mouthparts of caged
319
bees was due to cleaning and grooming behavior. Although we observed spores within the
320
foregut, separation of foregut and midgut during dissection may have contaminated the foregut.
321
Trophallaxis of honey bees has been shown to be a possible transmission pathway (Smith, 2012),
322
and here we demonstrated that the transmitted spores were most likely acquired from hygienic
323
behaviors, not from secretions from glands or regurgitation. Our results also suggest that
324
sanitation procedures similar to those used for N. apis in previous research (Bailey and Ball,
325
1991) could reduce transmission among individuals.
326
Examination of Malpighian tubules under light microscopy and with PCR produced the
327
same results- no observable infections but consistently positive PCR signals. We did not find a
328
method that prevented contact of Malpighian tubules with friable infected midgut tissues during
329
dissection. It is possible that germination of primary spores in the midgut could allow polar tubes
330
to breach the basal membrane and result in the placement of sporoplasms in tissues that are
331
appressed to the midgut (Solter and Maddox, 1998). While this could explain an occasional
332
finding of DNA in these tissues, it is highly unlikely that the sporoplasms could reach the
333
cephalic tissues. The finding of no DNA in hemolymph of heavily infected bees suggests that
334
the microsporidia probably escape the midgut tissues rarely if at all. The more intense infections
335
in caged bees compared to restrained bees cannot be explained by our data, but negative results
336
from more heavily infected caged bees that were dissected through the vertex rather than
337
mouthparts confirm that microsporidia DNA is not present in the cephalic tissues.
338
We did not observe microsporidian spores or vegetative forms in any tissues other than
339
the alimentary canal, even when PCR showed positive results for conventional dissections. We
340
noted that mature spores from ruptured midgut tissues apparently stick to the surface of other
341
dissected tissues and are difficult to remove by rinsing the tissues. Likewise, spores in the
342
mouthparts of caged bees were not easily removed. Our dissection and restraint methods were
343
designed to eliminate such contamination. qPCR results showing no DNA present for bees that
344
were sterilized and dissected via the head capsule vertex, and those that were individually
345
restrained to avoid social grooming do not support the hypothesis of infection in tissues other
346
than the midgut for either species. Our results demonstrated that N. ceranae has identical tissue
347
tropism with N. apis but overall spore production and timing of release from host midgut cells is
348
sufficiently different to provide a competitive advantage to N. ceranae.
349
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Bailey, L., Ball, B.V. 1991. Honey bee pathology. Academic Press, London ; New York. Becnel, J. J. 2012. Complementary Techniques: Preparations of Entomopathogens and Diseased Specimens for more Detailed Study using Microscopy. In "Manual of Techniques in Insect Pathology, Second Edition" [L.A. Lacey, Ed.] Academic Press, Great Britain. Becnel, J.J., Andreadis, T.G. 1999. Microsporidia in insects. In “The Microsporidia and Microsporidiosis” [M. Wittner, L. M. Weiss, Eds.] American Society for Microbiology Press, Washington, D.C. Chen, Y., Evans, J.D., Zhou, L., Boncristiani, H., Kimura, K., Xiao, T., Litkowski, A.M., Pettis, J.S. 2009. Asymmetrical coexistence of Nosema ceranae and Nosema apis in honey bees. J. Invertebr. Pathol. 101, 204-209. Chen, Y.P., Huang, Z.Y. 2010. Nosema ceranae, a newly identified pathogen of Apis mellifera in the USA and Asia. Apidologie. 41, 364-374. Chen, Y.W., Chung, W.P., Wang, C.H., Solter, L.F., Huang, W.F. 2012. Nosema ceranae infection intensity highly correlates with temperature. J. Invertebr. Pathol. 111, 264-267.
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Copley, T.R., Jabaji, S.H. 2012. Honeybee glands as possible infection reservoirs of Nosema ceranae and Nosema apis in naturally infected forager bees. J. Appl. Microbiol. 112, 15-24. Cordes, N., Huang, W.F., Strange, J.P., Cameron, S.A., Griswold, T.L., Lozier, J.D., Solter, L.F. 2012. Interspecific geographic distribution and variation of the pathogens Nosema bombi and Crithidia species in United States bumble bee populations. J. Invertebr. Pathol. 109, 209-216. De Graaf, D.C., Jacobs, F.J. 1991. Tissue-Specificity of Nosema-Apis. J Invertebr Pathol. 58, 277-278. Forsgren, E., Fries, I. 2010. Comparative virulence of Nosema ceranae and Nosema apis in individual European honey bees. Vet. Parasitol. 170, 212-7. Fries, I. 1993. Nosema-Apis - a Parasite in the Honey-Bee Colony. Bee World. 74, 5-19. Fries, I., Feng, F., daSilva, A., Slemenda, S.B., Pieniazek, N. J. 1996. Nosema ceranae n sp (Microspora, Nosematidae), morphological and molecular characterization of a microsporidian parasite of the Asian honey bee Apis cerana (Hymenoptera, Apidae). Europ J Protistol. 32, 356-365. Fries, I., Granados, R.R., Morse, R.A. 1992. Intracellular Germination of Spores of Nosema-Apis Z. Apidologie. 23, 61-70. Gisder, S., Hedtke, K., Mockel, N., Frielitz, M.C., Linde, A., Genersch, E. 2010. Five-year cohort study of Nosema spp. in Germany: does climate shape virulence and assertiveness of Nosema ceranae? Appl. Environ. Microbiol. 76, 3032-8. Higes, M., Garcia-Palencia, P., Martin-Hernandez, R., Meana, A. 2007. Experimental infection of Apis mellifera honeybees with Nosema ceranae (Microsporidia). J. Invertebr. Pathology. 94, 211-217. Higes, M., Martin-Hernandez, R., Meana, A. 2006. Nosema ceranae, a new microsporidian parasite in honeybees in Europe. J. Invertebr. Pathol. 92, 93-95. Huang, W. F., Bocquet, M., Lee, K.C., Sung, I.H., Jiang, J.H., Chen, Y.W., Wang, C.H. 2008. The comparison of rDNA spacer regions of Nosema ceranae isolates from different hosts and locations. J. Invertebr. Pathol. 97, 9-13. Huang, W. F., Jiang, J.H., Chen, Y.W., Wang, C.H. 2007. A Nosema ceranae isolate from the honeybee Apis mellifera. Apidologie. 38, 30-37. Klee, J., Besana, A.M., Genersch, E., Gisder, S., Nanetti, A., Tam, D.Q., Chinh, T.X., Puerta, F., Ruz, J.M., Kryger, P., Message, D., Hatjina, F., Korpela, S., Fries, I., Paxton, R. J. 2007. Widespread dispersal of the microsporidian Nosema ceranae, an emergent pathogen of the western honey bee, Apis mellifera. J. Invertebr. Pathol. 96, 1-10. Liu, T. P. 1992. Oocytes degeneration in the queen honey bee after infection by Nosema apis. Tissue Cell. 24, 131-8. Martin-Hernandez, R., Botias, C., Bailon, E.G., Martinez-Salvador, A., Prieto, L., Meana, A., Higes, M. 2012. Microsporidia infecting Apis mellifera: coexistence or competition. Is Nosema ceranae replacing Nosema apis? Environ. Microbiol. 14, 2127-2138. Martin-Hernandez, R., Meana, A., Garcia-Palencia, P., Marin, P., Botias, C., Garrido-Bailon, E., Barrios, L., Higes, M. 2009. Effect of Temperature on the Biotic Potential of Honeybee Microsporidia. Applied and Environ. Microbiol. 75, 2554-2557. Mcivor, C.A., Malone, L. A. 1995. Nosema-Bombi, a Microsporidian Pathogen of the Bumble Bee Bombus-Terrestris (L). New Zealand J. Zool. 22, 25-31. Paxton, R. J., Klee, J., Korpela, S, Fries, I., 2007. Nosema ceranae has infected Apis mellifera in Europe since at least 1998 and may be more virulent than Nosema apis. Apidologie. 38, 558-565. Solter, L.F., Becnel, J.J., Oi, D.H. 2012. Microsporidian entomopathogens. In “Insect Pathology, Second Edition. [F.E. Vega and H.K. Kaya, Eds.] Elsevier, San Diego. Solter, L. F., Maddox, J. V. 1998. Timing of an early sporulation sequence of microsporidia in the genus Vairimorpha (Microsporidia: Burenllidae). J. Invertebr. Pathol. 72, 323-329. Smith, M. L. 2012. The Honey Bee Parasite Nosema ceranae: Transmissible via Food Exchange? PLoS One. 7, e43319.
412 413
Figure 1
414
Inoculation of honey bees with microsporidia
415 416 417
Figure 2
418
Individually restrained bees (A) in microcentrifuge tube; (B) in feeding tube
419 420 421 422 423
Figure 3. Spore count in hindgut contents. (A) Nosema ceranae inoculations; (B) N. apis inoculations. The growth curves for mature spores were calculated by dividing the number of mature spores by the total spore count.
424
425 426
Table 1: qPCR results for infection detected in dissected tissues and rinsate from communally caged bees
Treatment
N. ceranae Conventional dissection
N. ceranae Dissected through vertex
N. apis Conventional dissection
N. apis Dissected through vertex
N. ceranae inoculation
Target tissue
Cta
No. spores
Std.Dev.
No. bees with detected infections (Nb)
Brain
NDc
ND
ND
0 (6)
Salivary gland
35.2
1743.0
2372.35
3 (6)
Hypopharyngeal gland
37.1
317.3
429.14
4 (6)
PBS rinse
35.5
1083.9
966.48
6 (6)
Brain
ND
ND
ND
0 (13)
Salivary gland
ND
ND
ND
0 (13)
Hypopharyngeal gland
ND
ND
ND
0 (13)
PBS rinse
ND
ND
ND
0 (13)
Brain
34.7
91.4
114.08
2 (4)
Salivary gland
35.6
39.2
44.29
2 (4)
Hypopharyngeal gland
34.0
104
--
1 (4)
PBS rinse
33.0
454.2
532.46
4 (4)
Brain
ND
ND
ND
0 (6)
Salivary gland
ND
ND
ND
0 (6)
Hypopharyngeal gland
ND
ND
ND
0 (6)
PBS rinse
ND
ND
ND
0 (6)
Oral cavity rinse
35.57
2009.5
1133.3
7 (7)
427 428
a
Ct = qPCR threshold cycle
429 430
b
Total number of bees examined
c
ND = not detected
Figure
Figure
Figure
431 432
Graphical abstract
433
Highlights:
434
436
Nosema ceranae produces more mature spores than Nosema apis in the honey bee host Nosema ceranae and Nosema apis develop only in honey bee midgut cells
437
One oral transmission pathway may be the result of hygienic behaviors
435
438 439
S0022-2011(13)00004-9 http://dx.doi.org/10.1016/j.jip.2013.01.001 YJIPA 6401
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Journal of Invertebrate Pathology
Received Date: Accepted Date:
16 October 2012 4 January 2013
Please cite this article as: Huang, W-F., Solter, L.F., Comparative development and tissue tropism in Nosema apis and Nosema ceranae, Journal of Invertebrate Pathology (2013), doi: http://dx.doi.org/10.1016/j.jip.2013.01.001
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1
Comparative development and tissue tropism in Nosema apis and Nosema ceranae
2 3 4 5
Wei-Fone Huang* and Leellen F. Solter
6 7
Illinois Natural History Survey, Prairie Research Institute, University of Illinois
8
1816 S. Oak Street, Champaign, IL 61801 USA
9 10 11 12 13 14 15
*Corresponding author: [email protected]
16 17
Abstract
18
The two etiological agents of nosema disease in honey bees, Nosema apis and Nosema ceranae
19
(Microsporidia: Nosematidae), reproduce in the midgut tissues of the host. Nosema apis is tissue
20
specific but the development and tissue tropism of Nosema ceranae is not well understood. Our
21
investigations compared development of the two phylogenetically related pathogens in all major
22
host tissues. Using microscopy, PCR and qPCR quantification to evaluate tissue tropism of
23
infected bees in communal cages and of individually restrained infected bees, we found no
24
detectable spores in cephalic or other body tissues except midgut tissues. Nosema DNA was
25
detected in Malpighian tubules but the tubules could not be separated from the alimentary tract
26
without release of spores from the midgut. Nosema DNA was not detected in hemolymph
27
sampled from the head capsule or the abdomen of infected bees. We confirmed that N. ceranae
28
only develops in midgut tissues. Spores of both species released from host midgut cells
29
accumulated in the hindgut lumen, and we noted differences in numbers and ratios of spore types
30
and in growth curves between the two pathogens. Nosema apis reached a consistent level of
31
spore production after 12 days post inoculation (dpi); Nosema ceranae spore production
32
increased linearly from 12-20 dpi and the number of mature Nosema ceranae spores was
33
consistently higher.
34 35 36
Keywords: Nosema; honey bee; tissue tropism; glands; transmission; development
37 38 39 40
Introduction
41
Apis mellifera, and cause chronic, debilitating disease. Infections frequently occur in high
42
prevalence in apiaries and deleteriously impact pollination services and production of honey
43
(Bailey and Ball, 1991; Fries, 1993). N. apis was the only known agent of honey bee nosemosis
44
until N. ceranae, a genetically related but distinct species, was described by Fries (1996) from
45
the Asiatic honey bee Apis cerana. N. ceranae was first reported infecting A. mellifera in Taiwan
46
in 2005 (Huang et al., 2007) and then in Europe (Higes et al., 2006), and was subsequently
47
identified in A. mellifera globally (Huang et al., 2008; Klee et al., 2007). The timing and means
48
of the invasion events are not known; N. ceranae was the predominant microsporidian pathogen
49
in A. mellifera by the mid-1990s in North America (Chen and Huang, 2010) and before 1990 in
50
Uruguay (Invernizzi et al. 2009).
51
Two microsporidian pathogens, Nosema apis and Nosema ceranae, infect the western honey bee,
Early morphological descriptions of N. apis were vague (McIvor and Malone, 1995), but
52
histological observations and tissue tropism studies were consistent: N. apis only infects the adult
53
stage of the host, and only reproduces in the midgut epithelium cells (Bailey and Ball, 1991; De
54
Graaf and Jacobs, 1991; Fries, 1993). The strict tissue tropism of N. apis is uncommon in the
55
genus. Most related Nosema species, including the closely related Nosema bombi infecting
56
bumble bees (McIvor and Malone, 1995; Fries et al., 2001), are systemic pathogens that
57
reproduce in the cells of most body tissues, including the gonads, and many species are
58
transmitted transovarially from infected females to their offspring (Becnel and Andreadis, 1999).
59
N. apis infection in queens may affect the development of oocytes (Liu, 1992) but the pathogen
60
is not transovarially transmitted (Webster et al. 2008).
61
N. ceranae, like N. apis, was described as a midgut pathogen of adult bees (Fries, 1996;
62
Fries 2010) and the two species are morphologically quite similar (Fries et al., 1996); therefore,
63
PCR has become the primary tool for differential diagnosis of honey bee nosematosis. When
64
PCR methods were used to detect N. ceranae, DNA signals were also observed in the cephalic
65
tissues (Chen et al., 2009; Copley and Jabaji, 2012; Gisder et al., 2010), but neither vegetative
66
forms nor mature spores have been observed in cephalic tissue smears and stains (Gisder et al.,
67
2010). Transmission electron microscopy showed changes in hypopharyngeal gland development
68
and cell micro-morphology in N. apis-inoculated bees but, like N. ceranae, no observable
69
infection (Liu, 1990). The possibility that N. ceranae infects hypopharyngeal and salivary glands
70
potentially changes the interpretation of transmission routes, pathogenesis, and host behavior,
71
and a reservoir function was suggested for the cephalic glands of both Nosema species (Copley
72
and Jabaji, 2012).
73
N. ceranae appears to have replaced or at least successfully competed with N. apis in
74
many, if not most, A. mellifera populations globally (Chen et al., 2009; Chen et al., 2012;
75
Martin-Hernandez, 2012). To address the characteristics of N. ceranae that have enabled it to
76
out-compete N. apis, an established pathogen, we compared the rate of development in the host,
77
production of spores, and tissue tropism. We used PCR and qPCR diagnoses for detection of N.
78
ceranae and N. apis in the tissues but rearing methods and dissection techniques were newly
79
designed to reduce the possibility that potentially uninfected target tissues were contaminated
80
with spores or DNA.
81 82
Materials and Methods
83
Nosema spp. isolates
84
Nosema ceranae was originally collected from campus apiaries at the University of Illinois at
85
Urbana-Champaign, and N. apis spores were provided by T. Webster at Kentucky State
86
University. To produce infective spores for experiments, honey bees were held in plastic cages
87
consisting of 480 ml HDPE lidded plastic cups with 3-mm hardware cloth tops (Webster et al.,
88
2004) and were inoculated with 105 spores of N. apis or N. ceranae. Bees were anesthetized on
89
ice, then fastened, dorsal side down, on foam boards with insect pins (Fig. 1). Upon warming,
90
bees were individually inoculated by allowing them to imbibe 2 µl sugar water containing 105 N.
91
apis or N. ceranae spores, a dosage that produces 100% infection (unpublished data). Bees were
92
returned to cages and at ≥10 days post inoculation (dpi), midgut tissues of infected bees were
93
excised, homogenized in a glass tissue grinder, and spores were isolated in sterile water
94
suspensions by centrifugation (Huang et al., 2007). Number and maturity of isolated spores were
95
checked under phase-contrast microscopy (400 X) using a Petroff-Hausser counting chamber.
96
Only freshly isolated spores were used in bioassays.
97
Inoculation of experimental bees
98
Newly emerged bees were harvested daily from a brood frame selected from a fumagillin-free
99
hive and were held in growth chambers at 34°C for 5 days. The bees were fed 50% sugar water
100
and pollen patties (MegaBee™, 15% pollen) ad libitum. Experimental bees were starved for 2 hr
101
and inoculated as described above. Only bees ingesting the full 2 µl suspension were included in
102
the trials. After allowing the bees to rest on the foam board for 30 min, they were transferred into
103
cages as described above, 30 bees per cage, and held in a 30°C growth chamber, 65% relative
104
humidity, 24 hr dark. The rearing temperature was reduced to 30°C for experiments because our
105
studies have shown that more intense infections develop at this temperature and high
106
temperatures may have adverse effects on infections (Martin-Hernandez et al., 2009; Chen et al.,
107
2012). In addition, our results are comparable to previous studies (Forsgren and Fries, 2010;
108
Paxton et al., 2007).
109
Examination of tissues
110
Honey bees were held for 14-15 days in communal cages as described above. Treated bees were
111
rinsed in 0.09% sodium hypochlorite (3% household bleach) with 0.2% tween20, then two rinses
112
in sterile water, and individually dissected. Tissues, including the midgut (with intense infection
113
levels), Malpighian tubules, fat body, crop and cephalic tissues, were excised and rinsed in PBS.
114
Small tissue samples were smeared on slides and examined under 400x phase contrast
115
microscopy. Giemsa stains were made of fresh tissue samples (Becnel, 2012) and were examined
116
under 400x bright field microscopy.
117
Spore counts
118
We observed spores in the hindgut contents of infected bees under phase-contrast microscopy
119
with refringency that was intermediate between the bright refractivity of mature “environmental”
120
(infective) spores and empty (germinated) spores (Solter et al., 2012). Based the morphology of
121
spores at specific time periods (Solter and Maddox, 1998; Higes, 2007; Solter et al., 2012) we
122
assumed that intermediately refringent spores with visible polar vacuoles are internally infective
123
“primary” spores (Fries et al., 1992; Solter et al. 2012). Other developing spores, e.g. immature
124
environmental spores, have similar refringency, although the polar vacuole is usually not visible.
125
Because it is even more difficult to distinguish spore stages and maturity based on refringency in
126
cell counting chambers (Petroff-Hausser), spores with similar refringency were assigned to the
127
same category. We thus counted three categories of spores, mature spores (high refringency),
128
primary/immature spores (low to medium refringency), and germinated spores (spore husks with
129
little refringency), every 2 days over the course of the test period. We conducted three trials; two
130
trials also served as inoculated positive control sets for our previous research (Huang et al.,
131
submitted).
132
Restraint of individual honey bees
133
We designed a restraint system to isolate test bees from colony members and eliminate grooming
134
and other hygienic behaviors. The tapered ends of 500-µl plastic microcentrifuge tubes were
135
removed, leaving an opening in the bottom of the tubes slightly larger than the diameter of the
136
head capsule but smaller than the thorax of A. mellifera. An anesthetized bee was placed head-
137
first into each tube, ensuring that the head emerged through the opening. Cotton was placed in
138
the remaining space to restrain the bee and the top was closed (Fig. 2A). After the bees warmed
139
and became active, they were inoculated with 105 N. ceranae spores in 2 µl water with a
140
micropipetter. After 30 min, the bees were transferred in the restraints to a 1.5 ml
141
microcentrifuge tube with 50% sugar water in the bottom that the bee could reach with its tongue
142
(Fig. 2B). The feeder was replaced every two days. The restrained bees were held in growth
143
chambers under the conditions described above and were dissected at 14 dpi to evaluate
144
infections in the midgut and cephalic tissues. We used either conventional dissection methods or
145
dissection through the head capsule vertex (see below) to dissect the restrained bees. The sugar
146
water removed from feeders 12 and 14 dpi was collected and diluted by adding ddH2O to remove
147
the sugars, to a total of 500 µl. The suspension was centrifuged and the supernatant was
148
discarded by pipetting, leaving approximately 10 µl of suspension. The dilution procedure was
149
repeated and the remaining 10-µl suspension was vortexed and observed under 400x phase
150
contrast microscopy, and DNA was extracted for PCR according to the following method.
151
Tissue sampling for PCR
152
Prior to surface sterilization of head capsules, the mouthparts of N. ceranae and N. apis
153
inoculated bees were individually sampled by pipetting 5 µl PBS several times between the
154
mandibles. DNA was then extracted from the rinsate. Both the communally caged and
155
individually restrained bees were sampled.
156
Bees were individually surface sterilized by washing them twice in cold 0.09% sodium
157
hypochorite with 0.2% tween20, and then rinsing twice in sterile water with 0.2% tween20. Bees
158
were dried on a clean paper towel and held on ice prior to dissection. Cephalic tissues, brain
159
(nerve tissues), hypopharyngeal glands and salivary glands of bees infected with either Nosema
160
species were collected, as well as foregut, midgut, Malpighian tubules, second tergum (with
161
attached tissues), and venom gland, of N. ceranae inoculated bees. Tissue smears were made
162
from small samples of fresh tissue and examined under 400x phase contrast microscopy. At least
163
one of the paired cephalic glands and approximately 2 mm3 cubic tissues of brain of each bee
164
were used for the DNA extractions.
165
The conventional method for excision of cephalic tissues we used was similar to that used
166
in published studies (Chen and Huang, 2010), and is briefly described as follows. The head was
167
removed with clean scissors and transferred to a Petri dish containing a few drops of PBS. The
168
cuticle was carefully cut through the oral cavity without touching internal tissues.
169
Hypopharyngeal glands, salivary glands and brain tissues were removed with clean forceps and
170
the tissues were washed twice in PBS. The tissues were transferred to a PCR tube containing
171
DNA extraction buffer. In addition to the tissues, 5 µl PBS (with no observable tissues) were
172
removed from the Petri dish and were added to DNA extraction buffer to determine if spores
173
were shed into the buffer. DNA was extracted for qPCR.
174
To prevent possible contamination with Nosema sp. spores during dissection, a second
175
method was used to excise cephalic tissues. Anesthetized, surface sterilized bees were dissected
176
without decapitation; the head was opened directly through the vertex area without touching the
177
mouthparts and pharynx. The cuticle was carefully removed, cutting along the vertical sutures
178
and sutures around the compound eyes, then tissues were removed from directly from the
179
opening with fine forceps. The tissues were placed in a few drops of PBS in a Petri dish and
180
separated under stereomicroscopy, then washed in PBS and transferred to DNA extraction buffer.
181
DNA was extracted as previously described. The PBS rinse was also sampled to check for shed
182
spores as previously described.
183
We sampled hemolymph from the head and abdomen of several infected surface
184
sterilized bees during dissection of tissues. Approximately 2 µl hemolymph was obtained from
185
the head capsule when dissecting via the vertex, and the same amount of abdominal hemolymph
186
was obtained via a razor cut of intersegmental membrane between the first and second abdominal
187
segments. DNA was extracted from hemolymph samples as described below.
188
DNA extraction
189
DNA was extracted using a modified Chelex method (Cordes et al., 2012). Dissected tissues,
190
approximately 2 mm3 per sample, were transferred into 50 µl Chelex buffer, 5% Chelex 100
191
(Biorad), 1ng/ul proteinase K and 5% tween20 in ddH2O, in a 200 µl PCR tube. The tubes were
192
vortexed at high speed for 15 s and centrifuged briefly to precipitate the Chelex buffer and tissue
193
to the bottom of the vial, then incubated using the following program: 56°C for 2 hr, followed by
194
95°C for 30 min, and a 4°C cool-down. The homogenate was centrifuged at 13,000 g for 10 min
195
and the supernatant retained for PCR.
196
qPCR and PCR
197
Quantitative PCR was performed using the SYBR green method. The specific primer set for N.
198
ceranae was Nc841f: GAG AGA ACG GTT TTT TGT TTG AGA; Nc980r: ATC CTT TCC
199
TTC CTA CAC TGA TTG, estimated amplicon size 140 bp. For N. apis, primers Na65f: CGT
200
ACT ATG TAC TGA AAG ATG GAC TGC and Na181r: AGG TCT CAC TCT TAC TGT
201
ACA TAT GTT AGC, estimated amplicon size of 116 bp, were designed using the SSUrRNA
202
sequence alignment. The reactions were carried out on an ABI 7900HT thermal cycler with
203
standard two-step PCR method using SYBR® Green PCR Master Mix (ABI). The annealing
204
temperature was 64.5C with 40 cycles. No cross-reaction between N. apis and N. ceranae or
205
non-specific PCR product was found. The results were analyzed using absolute quantification set
206
up in SDS 2.2 (ABI) with spore extractions as standards.
207
Spore extractions that were filtered, centrifuged, re-suspended, and counted were used as
208
standards. Four standard samples of both N. apis and N. ceranae in log dilutions of 5x10 to
209
5x104 spores were used to construct the standard curves for qPCR. We used identical methods
210
and volume for the standards and dissected tissues, and extracted the DNA at the same time. The
211
same primer sets were used for regular PCR diagnosis with the same reaction program, but with
212
Platinum® Taq (Invitrogen) following manufacturer’s directions.
213
Statistical analysis
214
The spore counts were analyzed by one-way ANOVA to determine significant differences using
215
SPSS software (IBM).
216 217
Results
218
Spores in hindgut lumen contents
219
Spore counts from the hindgut lumen contents are shown in Figure 3. Total numbers of the three
220
types of spores, mature, immature/primary, and germinated, were all significantly different
221
(P<0.001) between N. ceranae and N. apis over the course of the infection period. More mature
222
spores and fewer germinated and primary/immature spores were observed for N. ceranae than
223
for N. apis. We analyzed the ratio of types of spores in hindgut contents at 12 dpi and found 80-
224
95% mature N ceranae spores, while 57-83% of N. apis spores were mature. The ratio of N. apis
225
mature spores increased during the infection period (Fig. 3). The growth curve for germinated
226
spores was irregular in N. apis infections, but increased linearly to 16 dpi in N. ceranae
227
infections. The total number of primary/immature spores increased to 14 dpi and then dropped
228
rapidly for both N. apis and N. ceranae infections.
229 230
Microscopic and PCR evaluation of tissues from caged bees
231
Midgut spore counts exceeded 106 spores in all treated bees at 14 dpi for both N. ceranae and N.
232
apis. No spores or vegetative stages were observed in tissues other than the alimentary tract,
233
however, we observed spores attached to the surfaces of dissected tissues, including Malpighian
234
tubules and the tissues within the tergum. PCR diagnoses produced signals for most tissues but
235
the microsporidia did not appear to be developing within the cytoplasm of host cells. We noted
236
that heavily infected midgut tissues are extremely fragile and break down during dissection, but
237
microsporidia were not observed in the hemolymph extracted prior to the dissection of the hosts.
238
The efficiency of qPCR to quantify spores from DNA was in the range of 90-98% for N.
239
apis and 96-99% for N. ceranae. Six qPCR tests (three repeats for each test, three tests for each
240
sample) on different sample sets were performed to generate the results in Table 1. Standard
241
curve and PCR efficiencies were linear and consistent among the different reactions.
242
Quantitative PCR measurements for Nosema DNA in tissues were inconsistent for
243
infected communally caged honey bees that were dissected using the conventional method
244
(Table 1). Three different cephalic tissues, brain, salivary glands and hypopharyngeal glands
245
were compared, and no N. ceranae DNA was detected in brain tissues of six of the infected bees.
246
Because inconsistent DNA quantities were found in cephalic tissues among bees, we could not
247
evaluate differences among these tissues. Tissues from N. apis-inoculated bees produced similar
248
results, but we detected N. apis DNA in brain tissues of two of four inoculated bees. All PBS
249
rinsate remaining from the conventional dissections contained Nosema DNA. The rinsate from
250
external mouthparts, mandibles and partial labium contained several thousand spores based on
251
qPCR results.
252
There was no detectable N. ceranae or N. apis DNA in the cephalic tissues or in the PBS
253
used to rinse and separate tissues dissected through the head capsule vertex. The same group of
254
bees was used for both types of dissection, but an additional N. ceranae inoculation trial using
255
the vertex dissection method confirmed the results. There was no detectable Nosema DNA in the
256
hemolymph samples from caged bees.
257 258
Infections in restrained bees
259
We restrained and isolated bees only for N. ceranae inoculations. Mortality was higher for
260
individually restrained bees during the course of the experiment (75%), than for caged bees
261
(approximately 10%), most likely due to isolation and restriction to movement, and midgut spore
262
counts were lower. Caged bees fed the same inoculum had 5-37 million spores in midgut, while
263
restrained bees produced 1-3 million spores. Twelve restrained honey bees survived to 14 dpi.
264
Infections in caged bees were late stage and well-developed. Infections in restrained bees
265
consisted of predominantly mature environmental spores but the mortality rate suggests that bees
266
with extremely high levels of infection did not survive. No microsporidian stages were observed
267
in cephalic tissues under phase contrast microscopy or in qPCR diagnosis of these tissues that
268
were dissected from restrained bees using the conventional method or the dissection method via
269
the head capsule vertex. We found no spores in mouthpart rinses or sugar water at 12 and 14 dpi.
270 271
Discussion
272
We compared the development of infection based on spore accumulation in the hindgut, and
273
tissue tropism of the two related microsporidian pathogens infecting the western honey bee. In
274
previous studies, no spore types were distinguished and spores were counted in the midgut or in
275
the entire abdomen (midgut tissue and hindgut contents) (Forsgren and Fries, 2010; Martin-
276
Hernandez et al., 2009; Paxton et al., 2007), although both primary spores and empty
277
(germinated) spores can be observed under phase contrast microcopy (Solter et al., 2012). Most
278
bees we used for these experiments avoided defecating inside the cages, so the change in ratio of
279
the different spore types accumulated in the hindgut reflected the progress of infection in the
280
midgut. PCR-based diagnosis cannot distinguish primary, immature and mature environmental
281
spores. We noted a significantly lower ratio of primary/immature spores in the hindgut contents
282
of N. ceranae-infected bees than of N. apis-infected bees. Primary spores and, we assume,
283
immature environmental spores, do not survive outside the host and do not appear to be
284
transmissible among hosts, thus the higher ratio of environmental spores potentially provides N.
285
ceranae a competitive advantage. The lower primary/immature spore ratio in N. ceranae
286
infections could imply better suppression of host cell immune response, resulting in lysis of
287
midgut cells with more advanced infections and more mature spores than occurs in N. apis
288
infections. The ratio of environmental spores was high in both species after 14 dpi.
289
Our observations of mature environmental spores in host hindgut contents suggested that
290
proliferation rates differed for N. ceranae (approximately 8×106 spores/day), and N. apis
291
(approximately 4×106 spores/day) based on estimated linear growth curve slopes after the spore
292
count in the midgut tissues reached the plateau phase at 10-12 dpi (Forsgren and Fries, 2010;
293
Paxton et al., 2007). Spore accumulation in the hindgut trended higher for N. ceranae than N.
294
apis after 12 dpi on a per sample-day basis and was significantly higher overall. N. ceranae
295
spores appear to mature later in the infection process than do N. apis spores. This observation
296
corresponds to TEM observations of midgut infections (Higes et al., 2007).
297
Previous research detected DNA but no observable microsporidian stages in cephalic
298
tissues of N. apis infected bees, therefore we investigated possible cephalic infection by N.
299
ceranae using qPCR to evaluate infection development. We used different dissection and caging
300
methods (communal or isolated) to eliminate the possibility of contamination. DNA of both N.
301
apis and N. ceranae was detected by qPCR in cephalic tissues when conventional dissection
302
methods via the mouthparts were used, but excision of cephalic tissues directly though vertex of
303
the head capsule of both cage-reared and restrained bees eliminated touching of the mouthparts,
304
which could be a source of contamination in qPCR detections. No N. apis or N. ceranae DNA
305
was detected in these qPCR samples. Moreover, we found no detectable N. ceranae DNA in
306
hemolymph collected from the head or abdomen, suggesting the inability of N. ceranae to enter
307
the hemocoel as was also determined for N. apis using an antiserum technique (De Graaf and
308
Jacobs, 1991). Our results do not support the hypothesis that N. apis and N. ceranae infect the
309
honey bee cephalic glands and use them as reservoir tissues (Copley and Jabaji, 2012). Spores
310
and DNA of both N. apis and N. ceranae were found only in the alimentary tract.
311
The mouthparts of infected caged bees contained relatively high numbers of spores,
312
possibly due to hygienic grooming in cages holding multiple infected bees, some of which likely
313
defecated in the cages, releasing spores. Fecal matter not removed from mouthparts by surface
314
sterilization could, thus, be a source of contamination during conventional dissections of cephalic
315
tissues. Although we did not observe fecal material on cage surfaces, we did find spores of both
316
Nosema species in rinsate brushed from the cage walls. No spores were found in the mouthparts
317
or food source of individually restrained infected bees. Restrained bees were prevented from
318
performing hygienic behaviors and we suggest that presence of spores in mouthparts of caged
319
bees was due to cleaning and grooming behavior. Although we observed spores within the
320
foregut, separation of foregut and midgut during dissection may have contaminated the foregut.
321
Trophallaxis of honey bees has been shown to be a possible transmission pathway (Smith, 2012),
322
and here we demonstrated that the transmitted spores were most likely acquired from hygienic
323
behaviors, not from secretions from glands or regurgitation. Our results also suggest that
324
sanitation procedures similar to those used for N. apis in previous research (Bailey and Ball,
325
1991) could reduce transmission among individuals.
326
Examination of Malpighian tubules under light microscopy and with PCR produced the
327
same results- no observable infections but consistently positive PCR signals. We did not find a
328
method that prevented contact of Malpighian tubules with friable infected midgut tissues during
329
dissection. It is possible that germination of primary spores in the midgut could allow polar tubes
330
to breach the basal membrane and result in the placement of sporoplasms in tissues that are
331
appressed to the midgut (Solter and Maddox, 1998). While this could explain an occasional
332
finding of DNA in these tissues, it is highly unlikely that the sporoplasms could reach the
333
cephalic tissues. The finding of no DNA in hemolymph of heavily infected bees suggests that
334
the microsporidia probably escape the midgut tissues rarely if at all. The more intense infections
335
in caged bees compared to restrained bees cannot be explained by our data, but negative results
336
from more heavily infected caged bees that were dissected through the vertex rather than
337
mouthparts confirm that microsporidia DNA is not present in the cephalic tissues.
338
We did not observe microsporidian spores or vegetative forms in any tissues other than
339
the alimentary canal, even when PCR showed positive results for conventional dissections. We
340
noted that mature spores from ruptured midgut tissues apparently stick to the surface of other
341
dissected tissues and are difficult to remove by rinsing the tissues. Likewise, spores in the
342
mouthparts of caged bees were not easily removed. Our dissection and restraint methods were
343
designed to eliminate such contamination. qPCR results showing no DNA present for bees that
344
were sterilized and dissected via the head capsule vertex, and those that were individually
345
restrained to avoid social grooming do not support the hypothesis of infection in tissues other
346
than the midgut for either species. Our results demonstrated that N. ceranae has identical tissue
347
tropism with N. apis but overall spore production and timing of release from host midgut cells is
348
sufficiently different to provide a competitive advantage to N. ceranae.
349
References
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Bailey, L., Ball, B.V. 1991. Honey bee pathology. Academic Press, London ; New York. Becnel, J. J. 2012. Complementary Techniques: Preparations of Entomopathogens and Diseased Specimens for more Detailed Study using Microscopy. In "Manual of Techniques in Insect Pathology, Second Edition" [L.A. Lacey, Ed.] Academic Press, Great Britain. Becnel, J.J., Andreadis, T.G. 1999. Microsporidia in insects. In “The Microsporidia and Microsporidiosis” [M. Wittner, L. M. Weiss, Eds.] American Society for Microbiology Press, Washington, D.C. Chen, Y., Evans, J.D., Zhou, L., Boncristiani, H., Kimura, K., Xiao, T., Litkowski, A.M., Pettis, J.S. 2009. Asymmetrical coexistence of Nosema ceranae and Nosema apis in honey bees. J. Invertebr. Pathol. 101, 204-209. Chen, Y.P., Huang, Z.Y. 2010. Nosema ceranae, a newly identified pathogen of Apis mellifera in the USA and Asia. Apidologie. 41, 364-374. Chen, Y.W., Chung, W.P., Wang, C.H., Solter, L.F., Huang, W.F. 2012. Nosema ceranae infection intensity highly correlates with temperature. J. Invertebr. Pathol. 111, 264-267.
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Copley, T.R., Jabaji, S.H. 2012. Honeybee glands as possible infection reservoirs of Nosema ceranae and Nosema apis in naturally infected forager bees. J. Appl. Microbiol. 112, 15-24. Cordes, N., Huang, W.F., Strange, J.P., Cameron, S.A., Griswold, T.L., Lozier, J.D., Solter, L.F. 2012. Interspecific geographic distribution and variation of the pathogens Nosema bombi and Crithidia species in United States bumble bee populations. J. Invertebr. Pathol. 109, 209-216. De Graaf, D.C., Jacobs, F.J. 1991. Tissue-Specificity of Nosema-Apis. J Invertebr Pathol. 58, 277-278. Forsgren, E., Fries, I. 2010. Comparative virulence of Nosema ceranae and Nosema apis in individual European honey bees. Vet. Parasitol. 170, 212-7. Fries, I. 1993. Nosema-Apis - a Parasite in the Honey-Bee Colony. Bee World. 74, 5-19. Fries, I., Feng, F., daSilva, A., Slemenda, S.B., Pieniazek, N. J. 1996. Nosema ceranae n sp (Microspora, Nosematidae), morphological and molecular characterization of a microsporidian parasite of the Asian honey bee Apis cerana (Hymenoptera, Apidae). Europ J Protistol. 32, 356-365. Fries, I., Granados, R.R., Morse, R.A. 1992. Intracellular Germination of Spores of Nosema-Apis Z. Apidologie. 23, 61-70. Gisder, S., Hedtke, K., Mockel, N., Frielitz, M.C., Linde, A., Genersch, E. 2010. Five-year cohort study of Nosema spp. in Germany: does climate shape virulence and assertiveness of Nosema ceranae? Appl. Environ. Microbiol. 76, 3032-8. Higes, M., Garcia-Palencia, P., Martin-Hernandez, R., Meana, A. 2007. Experimental infection of Apis mellifera honeybees with Nosema ceranae (Microsporidia). J. Invertebr. Pathology. 94, 211-217. Higes, M., Martin-Hernandez, R., Meana, A. 2006. Nosema ceranae, a new microsporidian parasite in honeybees in Europe. J. Invertebr. Pathol. 92, 93-95. Huang, W. F., Bocquet, M., Lee, K.C., Sung, I.H., Jiang, J.H., Chen, Y.W., Wang, C.H. 2008. The comparison of rDNA spacer regions of Nosema ceranae isolates from different hosts and locations. J. Invertebr. Pathol. 97, 9-13. Huang, W. F., Jiang, J.H., Chen, Y.W., Wang, C.H. 2007. A Nosema ceranae isolate from the honeybee Apis mellifera. Apidologie. 38, 30-37. Klee, J., Besana, A.M., Genersch, E., Gisder, S., Nanetti, A., Tam, D.Q., Chinh, T.X., Puerta, F., Ruz, J.M., Kryger, P., Message, D., Hatjina, F., Korpela, S., Fries, I., Paxton, R. J. 2007. Widespread dispersal of the microsporidian Nosema ceranae, an emergent pathogen of the western honey bee, Apis mellifera. J. Invertebr. Pathol. 96, 1-10. Liu, T. P. 1992. Oocytes degeneration in the queen honey bee after infection by Nosema apis. Tissue Cell. 24, 131-8. Martin-Hernandez, R., Botias, C., Bailon, E.G., Martinez-Salvador, A., Prieto, L., Meana, A., Higes, M. 2012. Microsporidia infecting Apis mellifera: coexistence or competition. Is Nosema ceranae replacing Nosema apis? Environ. Microbiol. 14, 2127-2138. Martin-Hernandez, R., Meana, A., Garcia-Palencia, P., Marin, P., Botias, C., Garrido-Bailon, E., Barrios, L., Higes, M. 2009. Effect of Temperature on the Biotic Potential of Honeybee Microsporidia. Applied and Environ. Microbiol. 75, 2554-2557. Mcivor, C.A., Malone, L. A. 1995. Nosema-Bombi, a Microsporidian Pathogen of the Bumble Bee Bombus-Terrestris (L). New Zealand J. Zool. 22, 25-31. Paxton, R. J., Klee, J., Korpela, S, Fries, I., 2007. Nosema ceranae has infected Apis mellifera in Europe since at least 1998 and may be more virulent than Nosema apis. Apidologie. 38, 558-565. Solter, L.F., Becnel, J.J., Oi, D.H. 2012. Microsporidian entomopathogens. In “Insect Pathology, Second Edition. [F.E. Vega and H.K. Kaya, Eds.] Elsevier, San Diego. Solter, L. F., Maddox, J. V. 1998. Timing of an early sporulation sequence of microsporidia in the genus Vairimorpha (Microsporidia: Burenllidae). J. Invertebr. Pathol. 72, 323-329. Smith, M. L. 2012. The Honey Bee Parasite Nosema ceranae: Transmissible via Food Exchange? PLoS One. 7, e43319.
412 413
Figure 1
414
Inoculation of honey bees with microsporidia
415 416 417
Figure 2
418
Individually restrained bees (A) in microcentrifuge tube; (B) in feeding tube
419 420 421 422 423
Figure 3. Spore count in hindgut contents. (A) Nosema ceranae inoculations; (B) N. apis inoculations. The growth curves for mature spores were calculated by dividing the number of mature spores by the total spore count.
424
425 426
Table 1: qPCR results for infection detected in dissected tissues and rinsate from communally caged bees
Treatment
N. ceranae Conventional dissection
N. ceranae Dissected through vertex
N. apis Conventional dissection
N. apis Dissected through vertex
N. ceranae inoculation
Target tissue
Cta
No. spores
Std.Dev.
No. bees with detected infections (Nb)
Brain
NDc
ND
ND
0 (6)
Salivary gland
35.2
1743.0
2372.35
3 (6)
Hypopharyngeal gland
37.1
317.3
429.14
4 (6)
PBS rinse
35.5
1083.9
966.48
6 (6)
Brain
ND
ND
ND
0 (13)
Salivary gland
ND
ND
ND
0 (13)
Hypopharyngeal gland
ND
ND
ND
0 (13)
PBS rinse
ND
ND
ND
0 (13)
Brain
34.7
91.4
114.08
2 (4)
Salivary gland
35.6
39.2
44.29
2 (4)
Hypopharyngeal gland
34.0
104
--
1 (4)
PBS rinse
33.0
454.2
532.46
4 (4)
Brain
ND
ND
ND
0 (6)
Salivary gland
ND
ND
ND
0 (6)
Hypopharyngeal gland
ND
ND
ND
0 (6)
PBS rinse
ND
ND
ND
0 (6)
Oral cavity rinse
35.57
2009.5
1133.3
7 (7)
427 428
a
Ct = qPCR threshold cycle
429 430
b
Total number of bees examined
c
ND = not detected
Figure
Figure
Figure
431 432
Graphical abstract
433
Highlights:
434
436
Nosema ceranae produces more mature spores than Nosema apis in the honey bee host Nosema ceranae and Nosema apis develop only in honey bee midgut cells
437
One oral transmission pathway may be the result of hygienic behaviors
435
438 439