The queens of eusocial bees, ants and wasps mate only during
a very short period early in life and males therefore produce
ejaculates consisting of large numbers of high quality sperm. Such
extreme selection for high fecundity resulted in males investing
minimally into their somatic survival, including their immune system.
However, if susceptible males are unable to protect their reproductive
tissue from infections, they compromise queen fitness if they transfer
pathogens during mating. We used the honey bee Apis mellifera and investigated the course of infection of the sexually transmitted pathogen Nosema apis.
We predicted that honey bee males are susceptible but protect their
reproductive tissues from infections. We investigated the effects of N. apis
infections on the midgut, the accessory glands and the accessory testes
and quantified the consequences of infection on male survival and
fecundity. We found that N. apis is able to infect males, and as
infections progressed, it significantly impacted fertility and survival
in older males. Even though we confirm males to be able to minimize N. apis infections of their reproductive tissues, the parasite is present in ejaculates of older males. Consequently N. apis evolved alternative routes to successfully infect ejaculates and get sexually transmitted.
The mating biology of eusocial ants, bees and wasps is truly
spectacular because reproductive females (queens) only copulate during a
very brief period early in their life when they acquire and store a
lifetime’s supply of sperm1, 2, 3.
As queens never replenish sperm later in life, sperm use and economy
has been taken to inimitable extremes in those species that maintain
colonies with millions of workers3, 4, 5, 6 and/or survive in the field for decades4, 7, 8.
To provide queens with sufficient numbers of sperm, social insect males
evolved ejaculates that contain large numbers of sperm of exceptionally
high quality3, 9. The production and maintenance of such ejaculates prior to mating induces substantial costs to eusocial insect males5, 10.
They are therefore thought to operate at their physiological limits,
where even the slightest disturbance or alteration in their environment
can compromise their fertility11.
Additionally, to maximise their reproductive potential, these males
have been found to only minimally invest into their somatic survival5, 10.
The
immunity of social insect males is also affected by their genetics
because they originate from non-fertilized eggs and are haploid animals,
which has been hypothesized to additionally increase their
susceptibility to parasites12.
Therefore, it is particularly interesting to study immune challenges
resulting from parasitic infections in eusocial insect males. Empirical
work has confirmed that immune responses of social insect males are
consistently lower than those of female workers e.g. in leaf cutter ants13, wood ants14 or bumblebees15.
The resulting susceptibility of social insect males to parasitism is
thought to have resulted in a number of life history adaptations, which
reduce a male’s risk of getting infected: e.g. they have short life
spans, are completely dependent on workers and do not provide any help
for their colony. However, social insects are known to host a range of
parasites, which easily spread through their colonies consisting of
large numbers of related individuals including males16, 17.
Male susceptibility might therefore reduce a colony’s opportunity to
control infections and compromise colony fitness further, especially if
such infections reduce male fecundity, life expectancy or
competitiveness. Finally, male susceptibility could also allow pathogens
to spread through their bodies and eventually infect their reproductive
organs. In such a case, sexually transmitted pathogens could then
inflict cross-generational costs. However, sexually transmitted diseases
and their consequences have not been studied in great detail in social
insects18, 19, 20.
Here we hypothesize that social insect males should minimize the risk
of spreading a disease to queens by protecting their reproductive
tissues from parasite infections.
To test this idea, we used the honey bee A. mellifera ligustica and investigated the effects caused by an infection with the widespread fungal disease Nosema apis.
Honey bees make interesting study organisms to unravel effects of
infections on male immunity and fecundity for a number of reasons. First
they are known to host a wide variety of parasites21, 22
and second many of these are widespread and abundant enough among
colony members to pose a potential threat to males (drones).
Additionally, deformed wing virus was successfully identified in honey
bee ejaculates, suggesting that some honey bee diseases could be
sexually transmitted19, 23. More recently, N. apis
has been reported to be present in ejaculates of honey bee drones, and
artificial insemination experiments confirmed that this pathogen can in
principle be transmitted during mating24. Consequently, N. apis
is an ideal study organism to test the effects of parasites on male
fertility, and whether male susceptibility allows a disease to spread to
both the reproductive organs and the ejaculate. In order to test this,
we conducted a series of experiments to unravel the effects of a N. apis
infection on a male’s somatic tissue, by inspecting infected somatic
tissues of drones at different ages and comparing the survival of
infected and uninfected drones. We also investigated whether males are
able to protect their reproductive organs and their ejaculates from N. apis infections to minimize the risk of transmitting infective spores during copulation. To do this we measured the effects of a N. apis
infection on the drones’ reproductive organs at different ages, and
quantified the consequences of such an infection on their fecundity.
To collect N. apis spores, we sampled 20 foraging workers from five different N. apis
infected honey bee colonies housed at the University of Western
Australia and froze all the 100 bees we sampled for 2hrs at -20 °C. We
dissected out their midguts, pooled them in an Eppendorf tube containing
1 mL of DDI water and a 3 mm tungsten bead (Qiagen, Australia), and
homogenized them in a mixer mill (Retsch MM301, Australia) for 30 s at
25 Hz. Next, we layered 0.5 mL of the homogenate onto 1.5 mL of 100%
Percoll (Sigma-Aldrich, Australia) in a 2 mL Eppendorf tube, and
centrifuged it at 18,000 x g for 60 min at 4 °C, discarding the supernatant. The pellet containing the N. apis spores was resuspended using 1.5 mL DDI water. The sample was briefly vortexed and centrifuged again at 20,700 x g
for 5 min. We repeated this procedure three times before resuspending
the final pellet in 0.5 mL of DDI water to store at -80 °C. Prior to
infecting drones, we thawed the sample and diluted it with sugar syrup
to a final concentration of 10,000 spores/μL. Our previous work showed
that collecting N. apis spores as described above minimizes the effect on their viability25.
Drone breeding and infection
In
summer 2010, we bred drones in two different colonies according to
standard apicultural practices by providing each colony with one empty
drone comb. Two days prior to hatching, we placed the drone frames into
an incubator at 33 °C and 90% humidity and collected drones on the day
they emerged (day 0). The first 21 drones collected were used for the
following procedures: The faeces of three newly hatched drones was
inspected under a microscope for the presence of N. apis spores.
Next, we freeze-killed 6 newly hatched drones for 2 hours to dissect out
their midguts and microscopically check the gut tissue for evidence of a
N. apis infection. During the dissections we found that their midguts were translucent and detected no signs of a N. apis
infection, such as spores or morphological changes. In addition, we
freeze-killed 6 newly hatched drones to prepare histological samples by
embedding their sexual organs (accessory glands and testes) and inspect
them for signs of an infection with N. apis. Finally, we froze a further 6 newly hatched drones to determine the presence or absence of N. apis DNA using microsatellite markers. As we never detected any signs of N. apis in any of the 21 newly hatched males inspected (Table 1), we concluded that newly hatched males were not infected with N. apis, which is in accordance with the literature26.
Table 1: Number of honey bee males used to investigate the prevalence of Nosema apis in faeces, midgut, sexual organs and ejaculates of drones of 7 different ages.
All remaining eclosing drones were collected and each was fed with 1 μL (10,000 spores) of N. apis
spore solution (prepared as described above), before we returned them
to their maternal colonies. We recaptured 21 drones on each day 6, 9,
13, 15, 20, and 25 days post infection, and used them for the same
experiments as described above: 3 drones were immediately checked for
spores in their faeces, 6 drones were freeze-killed to dissect and
inspect their midguts, 6 drones were used for histological embedding and
inspection of their sexual organs, and 6 drones were checked for N. apis DNA using microsatellite markers. In addition, to investigate, whether sexually mature drones would be able to transmit N. apis
spores to the queen during mating, we microscopically inspected
ejaculates of 3 infected drones on days 13, 15, 20 and 25 days post
infection, respectively. An overview over the sample sizes used for each
experiment is provided in Table 1.
Histological confirmation of N. apis spore presence in reproductive tissue
After
collecting and freeze-killing 6 drones for each of the 7 age groups as
described above, we dissected their reproductive organs (accessory
glands and accessory testes). To avoid contamination, we cleaned all
equipment after each dissection with ethanol, bleach and water27.
Furthermore, we rinsed each tissue sample three times prior to
histological embedding in Hayes solution (0.15 M NaCl, 1.80 mM CaCl2, 2.68 mM KCl, 1.19 mM NaHCO3
, adjusted to pH 8.7 using NaOH, filtered (0.22 μm Millex® GP filter
unit, America). Next, we transferred each sample into a 2 mL tube and
covered it with 1.8 mL cold fixative (2.5% glutaraldehyde, 2%
paraformaldehyde) before placing it on a Stovall Belly Dancer TM
Shaker on 10 rpm for 30 min on ice at 4 °C overnight in a dark box. We
then transferred the samples into 0.1 M phosphate buffer at pH 7.4,
mixing them on ice for 15 min on a Belly Dancer on 10 rpm. Next we
placed them into 1% osmium tetroxide (Sigma-Alderich, Australia) for 2 h
before rinsing them in phosphate buffer. To dehydrate the tissue
samples, we first placed them in an ascending series of ethanol
concentrations (50, 70, 90, and 100%) followed by two washes in
propylene oxide and embedding into Epon Procure 812-Araldite resin
(Polyscience, Inc., Australia), using four steps of increasing
concentrations of propylene oxide to resin ratios (1:3, 1:1: 3:1 and
pure resin), all according to the manufacturer’s protocol. We finally
placed each tissue sample in moulds containing resin at 60 °C overnight.
All embedded organs were sectioned with glass knifes using an
ultramicrotome (LKB). The sections of 1-2 μm were then de-plasticized
using an aged, saturated solution of sodium hydroxide in 100% ethanol
(sodium ethoxide) for 5 min. Afterwards, we rinsed them in 100% ethanol
and water for 10 s each before placing them in 1% w/v hydrogen peroxide
for 7.5 min. Finally, we rinsed each section under tap water for 3 s and
stained them with Slidder’s Hematoxylin, Eosin and 1% w/v Biebrich
scarlet28.
Last we mounted the sections on microscopic slides using water-free
mounting media (Entellan®, Merkck Millipore, Australia) and investigated
them under an Olympus BX51 microscope (Olympus, Japan bright field,
Namarski optics with UPLAN FL objective lenses), taking digital photos
with an Olympus DP70 camera.
Detection of N. apis in male reproductive tissue using microsatellites
We used specific primers to test for the presence of N. apis
in the drones’ reproductive organs. To do this we dissected the
accessory glands and accessory testes from 6 drones per age group (see Table 1)
and rinsed them in Hayes solution. To extract DNA, we placed each
sample in 95% ethanol before transferring it into a vial with 500 μL
extraction buffer (0.1 M Tris, 0.05 M EDTA, 0.5 M NaCl, 1% w/v
Polyvinyl-pyrrolidone) and a 3 mm tungsten bead (Qiagen, Australia).
After homogenising each sample for 3 min in a mixer mill at 25 Hz, we
added 66 μL of 10% sodium dodecyl sulphate before centrifugation at
20,800 x g at 4 °C for 15 min. We added 445 μL of isopropanol to
600 μL of the supernatant and incubated the samples on ice for 15 min
before centrifugation at 20,800 x g at 4 °C for 15 min. We resuspended the DNA pellet in 500 μL of 70% ethanol and centrifuged each sample again at 20,800 x g
at 4 °C for 15 min. After discarding the supernatant, each pellet was
air dried for 20-45 min before being resuspended in 100 μL of DDI water
and centrifuged again at 20,800 x g at 4 °C for 15 min. We
collected the supernatant and determined the amount of DNA using a
NanoDrop (ND-1000 V3.2.1., America). Prior to PCR, we diluted each
sample to a final concentration of 50 ng DNA/ μL in DDI water.
We used N. apis primer sequences as published in the literature29,
purchased from Sigma-Alderich, Australia. The forward primer used was
5-GGGGGCATGTCTTTGACGTACTATGTA-3 and the reverse primer was
5-GGGGGGCGTTTAAAATGTGAAACAACTATG-3. We amplified DNA in a S1000™ Thermal
Cycler Chassis (Bio Rad, Australia). Each reaction contained 13.4 μL
sterile DDI water, 2 μL 10x Taq buffer, 0.1 μL Taq polymerase (Bio-Rad,
Australia, Cat# M0267X), 1.0 μL of each forward and reverse primer,
0.2 μL of 10% Triton-x100, 0.5 μL dNTP (Bio-Rad, Australia, Cat#
0447 L), and 2 μL of extracted DNA or sterile DDI water used as a
negative control. We denatured the DNA for 5 min at 94 °C (1x cycle);
which was followed by 30 reaction cycles consisting of denaturation for
15 s at 94 °C, primer annealing for 30 s at 61.8 °C, extension for 45 s
at 72 °C; and a final extension cycle of 7 min at 72 °C (1x cycle).
To confirm the absence or presence of N. apis
DNA in each sample, we ran 5 μL of the PCR amplification product on 1%
agarose gels (Promega, Australia,) in 1x Tris-borate-EDTA buffer with 2%
v/v of electrophoresis ethidium bromide (Merck, Australia). We used a
100 bp DNA ladder (Invitrogen, Australia, Cat# 15628-019) as a molecular
marker, and a N. apis spore sample as a positive control. For
gel electrophoresis we used a Bio Rad Mini-Sub® Cell GT Cell (Australia)
at 80 V, 400 mA for 30 min. We then photographed the gels in a Bio Rad
ChemiDoc™ XRS + System with Image Lab™ Software (Australia) and
determined the presence or absence of N. apis by checking each sample for species-specific gel bands.
The effect of N.apis infection on drone survival and sperm viability
In 2013, we conducted an additional experiment to quantify the effect of N. apis
on drone survival and sperm viability at different ages, and bred
drones in two different colonies as described above. After hatching, we
randomly allocated 20-30 drones to one of 25 cages (14 × 19.5 × 2.3 cm,
punctured metal sheet on one side and drone excluder on the other) and
placed them back into their maternal colonies for the next 24 h. We
recollected the cages and fed the drones with either 1 μL Nosema spore
solution (10,000 spores), or with 1 μL of 100% w/v sugar syrup as a
control. Consequently we ended up with 12 cages containing infected
drones and 14 cages containing uninfected drones. We resampled drones
12, 13, 16, 19, 20, 24 and 25 days post infection. For each cage, we
first counted all live and dead drones to quantify drone survival. We
anesthetized the surviving drones in chloroform to initiate ejaculation
and gently squeezed the drones’ abdomen between two fingers, until semen
appeared at the tip of the endophallus30. To quantify sperm viability, we used flow cytometry as described by Paynter et al. (2014)31.
In short, we collected around 2 μL of ejaculate per drone in 1 μL of
semen diluent (188.3 mM sodium chloride, 5.6 mM glucose, 574.1 nM
arginine, 684.0 nM lysine, 50 mM tris (hydroxymethyl) aminomethane, pH
8.7). We added 1 mL of semen diluent to each semen sample and gently
mixed it by turning the vial upside down until the ejaculate had fully
dispersed. To avoid mucus clogging up the capillary of the
flow-cytometer, we filtered a subsample of 200 μL through a 50 μm
diameter nylon mesh and added 800 μL of semen diluent to the filtered
sample. To differentially dye live and dead sperm, we used 400 μL of
sperm sample, added 2 μL of 1 mM SYBR 14 dye (Invitrogen, cat no.
L-7011) and incubated it in the dark for 10 min. We added 2 μL of 2.4 mM
Propidium Iodide (PI) (Invitrogen, cat no. L-7011) and incubated the
sample for 7 min in the dark. Sperm viability was quantified for a
minimum of 3000 sperm in a BD FACS Canto II digital flow cytometer
(America). The flow cytometer recorded SYBR 14 fluorescent emission in
the range 515–545 nm, and PI fluorescent emission in the range
670–735 nm without compensation for spectral overlap. We recorded height
rather than area of the voltage pulse generated by SYBR 14 and PI,
because honey bee sperm is exceptionally long (260 μm)1.
Sperm dyed with SYBR 14 were gated as live and those stained with PI
were gated as dead, using the FlowJo software package Version 7.6.5 for
Windows (TreeStar, USA). We used the autogate function of the Flowjo
software, except in 10 cases where the populations of live and dead
sperm could not be separated by the software. As is typically done in
these cases we therefore assigned the gates manually. We ignored doubly
stained cells, which occurred at very low frequencies. Statistical
analyses were performed using SPSS version 21 for Macintosh. Drone ages
were grouped for Figures, but statistical analyses were done using drone
age as a covariate.
We found that all drones fed with N. apis spores consequently developed an infection (Table 1),
which we were able to confirm by the presence of morphological changes
that became visible in the midgut of infected compared to non-infected
drones (Fig. 1).
We found that the midguts of newly hatched, non-infected drones
appeared transparent and remained translucent as drones matured (Fig. 1A,C).
In contrast, as the infected drones matured, their midguts developed
substantial swelling, lost their transparency and changed their colour
to a grey-white (Fig. 1B,D). Microscopic inspections confirmed that the epithelial cells of the midguts of infected drones were filled with N. apis spores (Fig. 2A-C) and we could observe large numbers of newly-released spores from burst cells (Fig. 2D).
Figure 1
Comparison of the midguts of mature male
honeybees that were either uninfected (A = gross morphology and
C = magnified) or infected with the microsporidian parasite Nosema apis (B = gross morphology and D = magnified).
Different views on the midgut and faeces of mature honeybee males infected with the microsporidian parasite Nosema apis:
A = midgut 20 days post infection, B = magnified view of an infected
midgut, C = close-up on the epithelial cells of an infected midgut.
D = faeces with pockets of spore masses (within the white border) among
other debris (brown).
When we compared dissected accessory glands and accessory testes of infected and non-infected drones (Fig. 3) we did not find any of the morphological signs of infection as described above for the midgut tissue (Fig. 1 & 2). Furthermore, we never found any N. apis
spores to be present in the sexual organs of infected drones of any
age, neither within the tissue nor in the lumen containing either sperm
or seminal fluid. This observation was confirmed histologically when
comparing the sexual tracts of non-infected and infected drones of
different ages. We found that the muscular tissue surrounding the
accessory gland and its epithelial cell layer gradually degenerated with
increasing drone age (Fig. 3), which was not the case for the muscular tissue surrounding the accessory testes (Fig. 3). However, neither the accessory glands nor the accessory testes tissues showed any visible signs of infection (Fig. 3).
Furthermore, despite a very careful microscopic inspection of both the
accessory testes and accessory glands, we never found a single N. apis spore in any of the infected males. However, our PCR analysis detected N. apis DNA in male reproductive tissues (Fig. 4).
This was the case for all drone age groups except for the newly emerged
drones, confirming that newly eclosed drones are not infected with N. apis. Finally, when we microscopically inspected ejaculated semen, N. apis spores were only present in drones aged 20 and 25 days (Fig. 5), but never in younger drones at 9, 13 or 15 days of age.
Figure 3
Longitudinal vertical cuts through the
accessory testes (top) and accessory gland (bottom) of male honey bees
at 0, 13 and 20 days post infection (p.i.) with the microsporidian
fungal parasite Nosema apis.
PCR amplification results showing DNA of the microsporidian parasite Nosema apis
in the reproductive tissues of infected, mature male honey bees.
Abbreviations: AG = accessory glands, AT = accessory testes,
Eja = ejaculate, -ve control = water used as negative control,+ve
control = samples known to contain Nosema apis DNA used as a positive control.
Self-ejaculated semen of a 20-day old honey bee male infected with the microsporidian parasite Nosema apis, showing sperm and mucus particles, as well as reproductive spores of the parasite.
Effect of N. apis infection on drone fecundity and survival
Sperm
viability data became available for 49 infected and 60 non-infected
males. We found that drone survival significantly decreased with
increasing age (ANCOVA, P < 0.001, see Table 2 for statistical details, Fig. 6 A).There was a significant effect of N. apis infection on male mortality, indicated by a significant treatment x age interaction (ANCOVA, df = 1, P = 0.013, Fig. 6 A).
Drone survival was similar between infected and non-infected drones
until they reached an age of about 16 days, after which mortality in
infected drones substantially increased compared to the control
treatment. Pairwise post-hoc t-tests using age groups revealed that a
significantly lower proportion of treated drones (20.7%) per cage
survived to the age of 24 to 25 days compared to control drones (54.7%),
with t = 3.5, df = 6 and p = 0.013. The results of the post-hoc tests
for the other age groups were non-significant.
Table 2: Survival of uninfected honey bee males and males infected with Nosema apis.
Comparison of honey bee males in different age groups that were either fed with spores of Nosema apis (Infected), or sugar syrup (Control): A)
Percentage of drone survival per cage, numbers in bars refer to the
number of cages used for each age group (20-30 bees per cage) B) Percentage of sperm viability per ejaculate. Numbers within bars indicate the number of drones used for each age group.
Similarly, sperm viability also decreased with male age (ANCOVA, P = 0.003, see Table 3 for statistical details, Fig. 6
B). We also found a significant treatment x age interaction term,
indicating that infected males lost their fertility faster than control
drones (ANCOVA, P = 0.013, see Table 3). The results of individual, pairwise post-hoc t-tests using age groups were not significant.
Table 3: Sperm viability of uninfected honey bee males and males infected with Nosema apis.
Our experiments revealed that N. apis is able to infect
drones, and that these infections built up to a point where they induced
significant costs for males. We found a reduction in fertility and life
span as drones aged (Fig. 6), as well as ejaculates of infected drones becoming contaminated with spores (Fig. 4).
Honey bee drones become sexually mature from 12 days after emergence
and consequently maintain their maximal fertility potential over a time
period of approximately 10 days32.
During that time, they participate in nuptial flights in order to find
and mate with virgin queens. The phenotypic expression of a N. apis
infection such as spores in the ejaculate and a reduction of drone
fertility and survival therefore affects these drones during their main
reproductive period. These findings indicate that drones which become
infected shortly after hatching will eventually face substantial fitness
costs and pose an infection threat to virgin queens in case they mate.
It would therefore be interesting to investigate, whether infected
drones indeed leave their colonies for nuptial flights and if so,
whether their mating success is compromised compared to that of
non-infected males. From a colony’s perspective, infected males might
choose not to participate in mating flights and mating in order not to
compromise the mating success of their non-infected brothers. Such
“altruistic” self-removal has been reported for honey bee workers after
prolonged CO2 narcosis or fed with the cytostatic drug
hydroxyurea. Both treatments increased worker-mortality, and surviving
foragers left their colonies, effectively committing altruistic suicide33. Such self-removal is also known from other social insects34. More research is required to quantify the risk of vertical transmission posed by males infected with N. apis.
Although our visual inspections, both morphologically as well as histologically, did not reveal any obvious signs of N. apis infections in reproductive tissues or their products, DNA of N. apis can be detected in both accessory glands and accessory testes. These findings indicate that N. apis
is able to establish low levels of infections in reproductive tissues
of drones. However, as we did not find spores in any of the reproductive
tissues we inspected, the pathogen seems unable to complete its
reproductive cycle or to build up an infection. Therefore, even though N. apis
is able to infect drone accessory testes and accessory glands, the
drones seem able to slow down or prevent this parasite from producing
spores within their reproductive tissue, thereby reducing the risk of
sexual transmission. This is an interesting finding, because N. apis
infections are already known to spread to different honey bee organs
such as the fat body, the malpigian tubules or the haemolymph35. In comparison, Nosema ceranae has been reported to infect drones at the pupal stage already36, to reduce drone body weight and life span and to induce physiological changes in honey bee queens37. Even though drones are more susceptible to N. ceranae than workers, they were are to develop higher tolerance to N. ceranae in specifically selected honey bee strains38.
It would therefore be interesting to study how males are able to
protect their reproductive tissue, and whether the costs associated with
suppressing N. apis infections result in the observed decrease in sperm viability and male survival.
We did find N. apis
spores in the semen of older drones, which raises the question of how
they were able to contaminate the ejaculate. Our dissection work
revealed that N. apis inflicts substantial damage to tissues such
as the midgut, which becomes increasingly more fragile with age and can
easily be damaged. Furthermore, N. apis is known to cause dysentery in honey bees, resulting in spore-containing faecal residues on bees39, 40.
It is therefore possible that the spores we detected in the ejaculate
of older drones are either caused by faecal contaminations of the
endophallus or haemorrhaging after tissue damage. Ejaculation is a
traumatic process as the drone contracts its abdominal muscles in order
to build up haemolymph-pressure, inducing the irreversible expulsion of
the endophallus and then the ejaculate. During a final step of
ejaculation, the tip of the drones’ endophallus bursts, breaks off and
is left inside the queen after copulation41,
causing the drones’ death. It therefore seems reasonable to assume that
infected tissues, which are more prone to damage and tear already,
release N. apis spores under the pressure build-up described
above, resulting in spore contamination of the ejaculate. Further
experimental work is required to unravel these proximate mechanisms.
However, if infections occur during the ejaculation process, a male’s
only counter measure to protect his ejaculate and mate would need to be
derived from immune traits present within the ejaculate. Interestingly,
drone ejaculates not only consist of sperm, but also of substantial
amounts of seminal fluid, produced by the accessory glands1, 42.
The latter is biochemically complex and contains a number of immune
proteins, some of them with well-known antifungal properties such as
chitinase43. Further research should therefore test the idea that seminal fluid is able to kill N. apis spores, in order to minimize the risk of N. apis establishing an infection inside the queen.
How to cite this article: Peng, Y. et al. Consequences of Nosema apis infection for male honey bees and their fertility. Sci. Rep.5, 10565; doi: 10.1038/srep10565 (2015).
Boomsma, J. J., Baer, B. & Heinze, J.The evolution of male traits in social insects. Annu. Rev. Entomol.50, 395–420, doi:10.1146/annurev.ento.50.071803.130416 (2005).
den Boer, S. P. A., Baer, B. & Boomsma, J. J.Seminal fluid mediates ejaculate competition in social insects. Science327, 1506–1509, doi:10.1126/science.1184709 (2010).
Sturup, M., Baer, B. & Boomsma, J. J.Short
independent lives and selection for maximal sperm survival make
investment in immune defences unprofitable for leaf-cutting ant males. Behav. Ecol. Sociobiol.68, 947–955, doi:10.1007/s00265-014-1707-x (2014).
Sturup, M., Baer-Imhoof, B., Nash, D. R., Boomsma, J. J. & Baer, B.When every sperm counts: factors affecting male fertility in the honeybee Apis mellifera. Behav. Ecol. Sociobiol.24, 1192–1198, doi:10.1093/beheco/art049 (2013).
O’Donnell, S. & Beshers, S. N.The role of male disease susceptibility in the evolution of haplodiploid insect societies. Proceedings Of The Royal Society B-Biological Sciences271, 979–983 (2004).
Baer, B., Krug, A., Boomsma, J. J. & Hughes, W. O. H.Examination of the immune responses of males and workers of the leaf-cutting ant Acromyrmex echinatior and the effect of infection. Insectes Sociaux52, 298–303 (2005).
Vainio, L., Hakkarainen, H., Rantala, M. J. & Sorvari, J.Individual variation in immune function in the ant Formica exsecta; effects of the nest, body size and sex. Evolutionary Ecology18, 75–84 (2004).
Gerloff, C. U., Ottmer, B. K. & Schmid Hempel, P.Effects of inbreeding on immune response and body size in a social insect, Bombus terrestris. Functional Ecology17, 582–589. (2003).
Knell, R. J. & Webberley, K. M.Sexually transmitted diseases of insects: distribution, evolution, ecology and host behaviour. Biological Reviews79, 557–581, doi:10.1017/S1464793103006365 (2004).
de Miranda, J. R. & Fries, I.Venereal and vertical transmission of deformed wing virus in honeybees (Apis mellifera L.). J. Invertebr. Pathol.98, 184–189, doi:10.1016/j.jip.2008.02.004 (2008).
Yue, C. & Genersch, E.RT-PCR analysis of deformed wing virus in honeybees (Apis mellifera) and mites (Varroa destructor). J. Gen. Virol.86, 3419–3424, doi:10.1099/vir.0.81401-0 (2005).
vanEngelsdorp, D. & Meixner, M. D.A historical review of managed honey bee populations in Europe and the United States and the factors that may affect them. J. Invertebr. Pathol.103, Supplement, S95 (2010).
Evans, J. D. & Schwarz, R. S.Bees brought to their knees: microbes affecting honey bee health. Trends Microbiol.19, 614–620, doi:10.1016/j.tim.2011.09.003 (2011).
Yue, C., Schroder, M., Gisder, S. & Genersch, E.Vertical-transmission routes for deformed wing virus of honeybees (Apis mellifera). J. Gen. Virol.88, 2329–2336, doi:10.1099/vir.0.83101-0 (2007).
Roberts, K.E.et al. The cost of promiscuity: sexual transmission of Nosema microsporidian parasites in polyandrous honey bees. Sci. Rep. 5, 10982, doi:10.1038/srep10982 (2015).
Peng, Y., Lee-Pullen, T. F., Heel, K., Millar, A. H. & Baer, B.Quantifying spore viability of the honey bee pathogen Nosema apis using flow cytometry. Cytometry Part A85, 454–462, doi:10.1002/cyto.a.22428 (2013).
Webster, T. C., Thacker, E. M., Pomper, K., Lowe, J. & Hunt, G.Nosema apis infection in honey bee (Apis mellifera) queens. Journal of Apicultural Research47, 53–57, doi:10.3896/IBRA.1.47.1.08 (2008).
Paynter, E.et al. Flow cytometry as a rapid and reliable method to quantify sperm viability in the honeybee Apis mellifera. 85, 472, doi:10.1002/cyto.a.22462 (2014).
Rueppell, O., Hayworth, M. K. & Ross, N. P.Altruistic self-removal of health-compromised honey bee workers from their hive. J. Evol. Biol.23, 1538–1546, doi:10.1111/j.1420-9101.2010.02022.x (2010).
Shorter, J. R. & Rueppell, O.A review on self-destructive defense behaviors in social insects. Insect. Soc.59, 1–10, doi:10.1007/s00040-011-0210-x (2012).
Webster, T. C., Pomper, K. W., Hunt, G., Thacker, E. M. & Jones, S. C.Nosema apis infection in worker and queen Apis mellifera. Apidologie35, 49–54, doi:10.1051/apido:2003063 (2004).
Traver, B. E. & Fell, R. D.Nosema ceranae in drone honey bees (Apis mellifera). J. Invertebr. Pathol.107, 234–236, doi:10.1016/j.jip.2011.05.016 (2011).
Huang, Q., Kryger, P., Le Conte, Y. & Moritz, R. F. A.Survival and immune response of drones of a Nosemosis tolerant honey bee strain towards N. ceranae infections. 109, 302 (2012).
Gochnauer, T. A., Furgala, B. & Shimanuki, H. in The hive and the honey bee: a new book on beekeeping which continues the tradition of “Langstroth on the hive and the honeybee” (ed Dadant and Sons) 615–662 (Dadant and Sons, 1975).
Baer, B., Zareie, R., Paynter, E., Poland, V. & Millar, A. H.Seminal fluid proteins differ in abundance between genetic lineages of honeybees. Journal of Proteomics75, 5646–5653, doi:10.1016/j.jprot.2012.08.002 (2012).
Baer, B., Heazlewood, J. L., Taylor, N. L., Eubel, H. & Millar, A. H.The seminal fluid proteome of the honeybee Apis mellifera. Proteomics9, 2085–2097, doi:10.1002/pmic.200800708 (2009).
The authors acknowledge the facilities, and the scientific and
technical assistance of the Australian Microscopy & Microanalysis
Research Facility at the Centre for Microscopy, Characterisation &
Analysis, The University of Western Australia, a facility funded by the
University, State and Commonwealth Governments. We also sincerely thank
the following people working at the University of Western Australia:
Thomas Stewart and Michael Archer for their assistance with histology,
Elke Ströher and Tiffane Bates for their technical assistance and Julia
Grassl for her intellectual input. This work was funded through two
Future Fellowships offered by the Australian Research Council (to B.B.
and A.H.M) and a ARC Linkage Project (to B.B. and A.H.M). The Rural
Industries Research and Development Corporation provided a PhD
scholarship to support Yan Peng.
Centre for Integrative Bee Research (CIBER), Bayliss Building
M316, The University of Western Australia, Crawley, WA 6009, Australia
Yan Peng,
Barbara Baer-Imhoof &
Boris Baer
ARC Centre of Excellence in Plant Energy Biology, Bayliss
Building M316, The University of Western Australia, Crawley, WA 6009,
Australia
A. Harvey Millar
Contributions
Y.P. and B.B.I. conducted the experiments, analysed the data and
contributed towards the writing of the paper, A.H.M. and B.B. supervised
the work, analysed the data and wrote the paper. All authors reviewed
the manuscript and agreed to its submission.
Competing financial interests
The authors declare no competing financial interests.
PROFESOR UNIVERSITARIO DICE: LA ÚNICA RAZÓN POR LA QUE MUEREN LAS ABEJAS ES LA VARROA
Abajo traducido a español
A
lifelong beekeeper and Mississippi State University Extension Service
apiculture specialist offers an unusual list of reasons for bee colony
death.
“My
top three reasons for bee colony death are Varroa mites, Varroa mites
and Varroa mites,” said bee expert Jeff Harris. “This is my sarcastic
response to the heavy emphasis in the press on the effects of
insecticides and other pesticides on honey bees.
Varroa
mites -- such as this one attached to a honeybee -- transmit viruses,
weaken bee health and factor prominently in the decline of bee
populations. (Photo by USDA-ARS/Steve Ausmus)
“Please
don’t misunderstand me. Insecticides and other pesticides kill honey
bees, either acutely by direct exposure to the chemicals or as part of a
group of stressors that kills honeybees,” he said.
But,
Harris said, there is no conclusive link between insecticide or
pesticide use and the widespread deaths of honey bee colonies that have
been occurring in the U.S., Canada and parts of Europe.
“What
is lost by an oversimplified view of colony health is that honey bees
suffer from myriad parasites, diseases and other stressors that are more
commonly associated with the death of the colony,” he said. “Most
scientists studying honey bees would rank Varroa mites and the viruses
they vector to honey bees as, hands down, the number one killer of honey
bees in the world. Most non-beekeeper members of the public have never
heard of Varroa mites. Even some new beekeepers don’t know what they
are.”
Varroa
mites are external parasites that lay eggs in the brood cells within
the hive and emerge attached to the host when the bee hatches out of its
cell.
“Imagine
a tick the size of a basketball attached to your neck,” Harris said.
“Varroa mites attach to honeybees and suck their hemolymph, which is
similar to blood in humans.”
Varroa mites also transmit diseases to honey bees. Harris estimated mites vector about 18 different viruses.
“Varroa
mites reproduce rapidly and reduce the health of the colony to the
point the colonies fail, or collapse,” Harris said. “We have found
colonies with ample stores of honey and either no bees or a handful of
bees left in the hive. As scientists, we had no doubt: high mite
populations vector high levels of viruses to honey bees that will
ultimately kill the colony.”
Beginning
in 2006, when episodes of high colony mortality were first reported,
millions of dollars have been spent on research into the causes of what
became known as Colony Collapse Disorder.
“Scientists
came to the conclusion that multiple factors cause these unusually high
death rates of bee colonies in some commercial operations,” Harris
said. “It also became apparent that different sets of stressors cause
losses for different beekeepers.”
Some
beekeepers lost colonies because of a combination of inadequate
nutrition related to periods of agricultural drought, stress related to
honey bee transportation, and parasitism by Varroa mites.
“Although
insecticides were acknowledged as contributing to the demise of bee
colonies, in most of the key studies into the causes of Colony Collapse
Disorder, scientists emphasized the factors causing the most significant
problems for honey bees were Varroa mites and the viruses they transmit
to honey bees,” he said.
At
first, the primary method for treating Varroa mites was insecticide,
but some mite populations became genetically resistant to the
insecticides. Other treatment options with limited effectiveness involve
mechanical methods, such as drone-brood trapping or freezing, or
natural methods, such as dusting colonies with powdered sugar to
increase the bees’ grooming behaviors, which results in mite removal.
“One
extremely bright glimmer of hope in the battle against Varroa mites is
the selective breeding of lines of honey bees that exhibit strong mite
resistance,” said Audrey Sheridan, entomology research and Extension
associate with the Mississippi Agricultural and Forestry Experiment
Station. Sheridan said Harris has brought to MSU his extensive bee
breeding experience from his former employment at the U.S. Department of
Agriculture Bee Lab in Baton Rouge
Beginning
in 1997, Harris and his USDA colleagues selected for bees that have a
trait termed Varroa Sensitive Hygiene. Bees with the trait can detect
Varroa mites in the combs of their nests, and they remove the bee pupae
infested by the mites.
This nest-cleaning behavior stops the mites’ reproductive cycle.
“Jeff
is working to improve stocks of VSH bees specifically for Mississippi’s
beekeeping environment,” she said. “The big take-home message from
scientific research is that our biggest single health issue in
beekeeping can be mitigated by using stocks of bees bred for resistance
to a parasite.
“We
just need to get people to try and use these important lines of bees
and do what they can as beekeepers to keep mite populations under
control,” she said.
En
Virtud de La Opinión del Profesor Asistente de Investigación en
Entomología Jeffrey Harris La Unica Razón de la muerte de las abejas es
la varroa. Hizo ONU article in this 2012 sobre tema
Un apicultor de toda la vida y el Servicio de Extensión de la
Universidad Estatal de Mississippi especialista en apicultura ofrece una
lista inusual de razones de la muerte de colonias de abejas.
"Mis tres principales razones de la muerte de colonias de abejas son
los ácaros varroa, ácaros Varroa y ácaros Varroa", dijo el experto en
abejas Jeff Harris.
"Esta es mi respuesta sarcástica a la fuerte énfasis en la prensa sobre
los efectos de los insecticidas y otros pesticidas en las abejas de
miel.
Ácaros Varroa - como éste adjunta a una abeja - transmitir virus,
debilitar salud de las abejas y el factor de forma destacada en la
disminución de las poblaciones de abejas. (Foto por el USDA-ARS / Steve Ausmus)
"Por favor, no me malinterpreten.
Los insecticidas y otros pesticidas matan a las abejas de miel, ya sea
de forma aguda por la exposición directa a los productos químicos o como
parte de un grupo de factores de estrés que mata abejas ", dijo.
Pero, Harris dijo, no existe una relación concluyente entre insecticida
o el uso de plaguicidas y las muertes generalizadas de las colonias de
abejas de miel que han estado ocurriendo en los EE.UU., Canadá y partes
de Europa.
"Lo que se pierde por una visión simplista de la salud de las colonias
es que las abejas sufren de parásitos, enfermedades y una miríada de
otros factores de estrés que están más comúnmente asociados con la
muerte de la colonia", dijo.
"La mayoría de los científicos que estudian las abejas podrían
clasificar a los ácaros Varroa y los virus que vector para las abejas
como, las manos hacia abajo, el asesino número uno de las abejas de miel
en el mundo. La mayoría de los miembros no apicultor del público nunca han oído hablar de los ácaros Varroa. Incluso algunos de los nuevos apicultores no saben lo que son ".
Ácaros Varroa son parásitos externos que ponen huevos en las celdas de
cría dentro de la colmena y emergen adjuntan al host cuando las
escotillas de abejas fuera de su celda.
"Imagina una garrapata del tamaño de una pelota de baloncesto conectado a su cuello", dijo Harris. "Ácaros Varroa se adhieren a las abejas y chupar su hemolinfa, que es similar a la sangre en los seres humanos."
Ácaros Varroa también transmiten enfermedades a las abejas de miel. Harris estima ácaros del vector de aproximadamente 18 virus diferentes.
"Ácaros Varroa se reproducen rápidamente y reducir la salud de la
colonia hasta el punto de las colonias fallan, o colapso", dijo Harris.
"Hemos encontrado colonias con amplios almacenes de miel y, o bien no
hay abejas o un puñado de abejas que quedan en la colmena.
Como científicos, teníamos ninguna duda: las poblaciones altas de
ácaros vector altos niveles de virus en las abejas de miel que en última
instancia va a matar la colonia ".
A partir de 2006, cuando se informó por primera episodios de mortalidad
elevada colonia, millones de dólares se han gastado en la investigación
de las causas de lo que se conoció como despoblamiento masivo.
"Los científicos llegaron a la conclusión de que varios factores causan
estos inusualmente altas tasas de mortalidad de las colonias de abejas
en algunas operaciones comerciales", dijo Harris. "También se hizo evidente que los diferentes conjuntos de factores de estrés causan pérdidas para diferentes apicultores."
Algunos apicultores perdieron las colonias debido a una combinación de
una nutrición inadecuada en relación con los períodos de sequía
agrícola, el estrés relacionado con el transporte de abejas de miel, y
el parasitismo de los ácaros Varroa.
"Aunque los insecticidas fueron reconocidos como contribuir a la
desaparición de las colonias de abejas, en la mayoría de los estudios
clave sobre las causas del desorden del colapso de colonias, los
científicos hicieron hincapié en los factores que causan los problemas
más significativos para las abejas eran los ácaros Varroa y los virus
que transmiten a la miel abejas ", dijo.
Al principio, el método principal para el tratamiento de los ácaros
Varroa fue insecticida, pero algunas poblaciones de ácaros se convirtió
genéticamente resistentes a los insecticidas.
Otras opciones de tratamiento con eficacia limitada implican métodos
mecánicos, como la captura o la congelación-drone cría, o los métodos
naturales, como colonias espolvorear con azúcar en polvo para aumentar
las conductas de aseo de las abejas, que se traduce en la eliminación de
ácaros.
"Un destello muy brillante de esperanza en la lucha contra la varroasis
es la cría selectiva de líneas de abejas que exhiben una fuerte
resistencia del ácaro", dijo Audrey Sheridan, investigación entomología y
Extensión asociada con la Estación Experimental Agrícola y Forestal de
Mississippi.
Sheridan dijo Harris ha traído a MSU su amplia experiencia de cría de
abejas de su antiguo empleo en el Departamento de Agricultura de los Bee
Lab en Baton Rouge
A partir de 1997, Harris y sus colegas del USDA seleccionados para las
abejas que tienen un rasgo denominado Varroa Sensible Higiene.
Abejas con el rasgo pueden detectar los ácaros Varroa en los panales de
sus nidos, y eliminar las pupas de abejas infestadas por los ácaros.
Este comportamiento de limpieza nido detiene ciclo reproductivo de los ácaros.
"Jeff está trabajando para mejorar las poblaciones de abejas VSH
específicamente para el entorno de la apicultura del Mississippi", dijo.
"El gran mensaje final de la investigación científica es que nuestro
mayor problema de salud individual en la apicultura puede ser mitigado
mediante el uso de las existencias de las abejas criadas para la
resistencia a un parásito.
"Sólo tenemos que hacer que la gente para tratar de utilizar estas
líneas importantes de abejas y hacen lo que pueden como apicultores para
mantener las poblaciones de ácaros bajo control", dijo.
Apicultores zamoranos y gallegos promueven una asociación interregional
Los productores pretenden fomentar la formación y divulgar la importancia de la actividad para la polinización de las plantas
A. SAAVEDRA
Los apicultores de las comarcas de zamoranas de Sayago, Sanabria,
Carballeda y Los Valles promueven la creación de una asociación
transfronteriza, "Zánganos Abelleiros", junto con los apicultores de
Orense, Tras Os Montes y posiblemente de León. Más de un centenar de
apicultores participaron la pasada semana en el homenaje dedicado a uno
de los maestros de la apicultura, Román Cid Álvarez, en Celanova donde
se trató la posibilidad de asociarse. La mayor parte de los promotores
de esta agrupación son apicultores alumnos del orensano Román, que ha
dedicado más de 30 años a las abejas y a enseñar a otros aficionados y
profesionales. Los alumnos le reconocen además de su conocimiento, el
ser un profesional colaborador y participativo, dispuesto a prestar su
conocimiento y experiencia, además de dar respuesta a los problemas que
surgen en las explotaciones.
La futura agrupación centrará su
actividad en divulgar la apicultura y su importancia en la polinización
de las diferentes especies vegetales, a parte de la producción de miel.
En estos momentos uno de los problemas importantes es la ausencia de
enjambres naturales que prácticamente mueren en un 100 si no se tratan
contra la enfermedad de la varroa, como señalan los apicultores de
Sanabria y Carballeda. Otro problema incipiente es la detección de
avispas asiáticas, Vespa velutina, en la provincia de Orense, limítrofe
con la comarca de Sanabria y las tierras lusas del norte de Tras Os
Montes.
Entre 110-120 apicultores de la zona limítrofe no
descartan crear la agrupación e incluir profesionales de León junto a
Orense, Zamora y tras Os Montes. Entre las propuestas de actividades que
se barajan están las de hacer cursos especializados y divulgar el mundo
de las abejas entre entre niños y adultos.
En estos momentos
los apicultores estiman que este año puede haber una buena cosecha de
miel en Sanabria y Carballeda, aunque en el invierno ha habido muchas
bajas. La floración de la primavera y el calor augura uno de los buenos
años en toda la zona, pendiente de evaluar si ha habido daños por las
tormentas. El sector ha recibido con preocupación el interés de la Junta
de Castilla y León en defender a los apicultores trashumantes en
perjuicio de los apicultores estantes asentados en las comarcas
zamoranas, en contraposición a la política de la Xunta de Galicia que ha
limitado los asentamientos trashumantes en su territorio.
