Transovarial transmission of Yersinia pestis in its flea vector, Xenopsylla cheopis

doi:10.21203/rs.3.rs-3397969/v1

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Transovarial transmission of Yersinia pestis in its flea vector, Xenopsylla cheopis

https://doi.org/10.21203/rs.3.rs-3397969/v1

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Yersinia pestis is the causative agent of bubonic plague, a deadly flea-borne disease responsible for three historic pandemics. Today annual cases of human disease occur worldwide following exposure to Y. pestis infected fleas that can be found within the rodent population where plague activity cycles between epizootic outbreaks and extended periods of apparent quiescence. Flea transmission of Y. pestis is most efficient in “blocked” fleas that are unable to feed, whereas mammalian transmission to fleas requires a susceptible host with end-stage high titer bacteremia. These facts suggest alternative mechanisms of transmission must exist to support the persistence of Y. pestis between epizootic outbreaks. In this work, we addressed whether vertical transmission could be a mechanism for persistent low-infection across generations of fleas. We demonstrate that Y. pestis infection of the Oriental rat flea, Xenopyslla cheopis, spreads to the reproductive tissues and is found in eggs produced by infected adult fleas. We further show that vertical transmission of Y. pestis from eggs to adults results in midgut colonization indicating a strong probability that it can reenter the sylvatic plague cycle.

Biological sciences/Microbiology/Pathogens

Health sciences/Diseases/Infectious diseases/Bacterial infection

flea

Xenopsylla

Yersinia

plague

transovarial transmission

Risk of plague, caused by Yersinia pestis, exists in endemic areas present in four continents due to the stable transmission cycle between rodents and their associated fleas ¹. Plague cycles between quiescent phases that may last decades and explosive epizootic outbreaks that increase human exposure ^2-6. Acquisition and regurgitative transmission of Y. pestis by fleas are inefficient mechanisms, associated with a shortened lifespan of the flea or rodent host ^7,8. Collectively, these features are seemingly incompatible, indicating that, despite over 100 years of intensive research on flea transmission of plague, there remain significant gaps in our understanding of the vector-animal cycle of Y. pestis. Herein, we found that vertical transmission is a missing gap in knowledge which can account for sustained Y. pestis infection of a flea population. We demonstrate that laboratory-reared and infected plague vector, Xenopsylla cheopis, transmits viable Y. pestis from adults to eggs that is subsequently successfully passed through all life stages. Further evidence indicates that transovarial transmission occurs throughout the life cycle of the infected flea. Together these data implicate transovarial transmission as a mechanism capable of sustaining Y. pestis in the environment for extended periods without detectable plague activity.

Yersinia pestis is the causative agent of plague, which is classified as a prioritized re-emerging zoonotic disease within the United States. Worldwide plague foci persist due to enzootic sylvatic cycles consisting of burrowing rodents and their respective flea vectors, which are a crucial component for transmission ^9–15. Numerous flea species and primarily rodent hosts are implicated in maintaining the sylvatic plague cycle, with mice, rats, prairie dogs, and ground squirrels commonly found in established plague foci ^16,17. Fleas can transmit Y. pestis as early as day 1 post-infection, in a poorly understood mechanism involving regurgitation from the midgut with an enhanced efficacy that results from high flea density ¹⁸. Once in the midgut, bacteria respond to environmental cues that activate expression and production of an exopolysaccharide matrix, which develops into an infectious biofilm ^19,20. During feeding the biofilm localized to the proventriculus occludes ingestion and causes regurgitation which deposits Y. pestis in the dermis of the human or animal host. From this site, as few as one bacterium is sufficient to cause lethal bubonic plague.

Early studies of the rodent-flea plague cycle documented proventricular blockage and its potential correlation to transmission in fleas known to vector Y. pestis. Blockage and transmission by blocked fleas was most frequent in the rat flea, Xenopsylla cheopis, with more than 50% occurrence, whereas blockage was very infrequent in other plague vectors including Oropsylla montana ²¹. Recent work demonstrated that occurrence of proventricular blockage in Y. pestis-infected O. montana could be enhanced by using rat blood feeding compared to mouse blood ²². This observation suggests that increased populations of rats should increase the likelihood of epizootic plague, however this is not observed in naturally occurring plague foci. In addition to biofilm-mediated proventricular blockage, Y. pestis can be transmitted through a regurgitative mechanism shortly after the flea becomes infected ²³. This mechanism appears to be independent of blood meal source, but can also result in non-productive transmission suggesting it may slow epizootic spread of disease ²⁴. Although these two methods of transmission are believed to require distinct proteins from the bacteria and fleas, both occur from the midgut. There have been no reports of Y. pestis localization to salivary glands, and no indication that bacteria are able to escape the midgut of fleas. Thus, the collective data led to the prevailing model for Y. pestis transmission involving continuous cycling between adult fleas and susceptible rodent hosts. With these parameters, the persistence of Y. pestis in the environment where there is little to no plague activity cannot be modeled.

In this work, we therefore addressed the historic paradigm that Y. pestis is confined to the midgut and alimentary canal in fleas with the underlying hypothesis that vertical transmission in the flea could account for long periods of reduced plague activity in a given endemic area. Using a highly sensitive fluorescent reporter with confocal microscopy and additional imaging with transmission electron microscopy, we reexamined the distribution of Y. pestis following membrane feeding of X. cheopis in male and female fleas and their progeny.

Yersinia pestis escapes to reproductive tissue in the flea vector Xenopsylla cheopis.

To understand Y. pestis interactions in fleas, we used a highly sensitive expression system for the fluorescent reporter tdTomato as a means to localize bacteria following infection of X. cheopis fleas in an artificial membrane feeder. With this approach, we established the behavior of fluorescent Y. pestis in the midgut using confocal microscopy and followed an apparent Y. pestis-containing biomass that developed and peaked in the midgut on day 3 post-infection (Fig. 1A-D). In some of the fleas collected mainly on day 3, brightly fluorescing Y. pestis could be seen in the esophagus, similar to previous reports (representative shown in Fig. 1C) ^22,25. By day 7, the Y. pestis containing biomass decreased, consistent with previous studies (Fig. 1D-E). The kinetics of the biomass suggests correlation with competency for early phase transmission. In parallel, bacterial load was quantified in pools of 3 midguts per group, demonstrating similar median values for colony forming units (CFU) across the 7-day time course (Fig. 1F). These data establish that while the kinetics of the Y. pestis-containing biomass change during the 7-day course of infection, viable bacterial load is relatively constant in the flea midgut.

To determine whether Y. pestis may be localized outside of the midgut, we examined the ovarioles and the testes of fleas dissected from the various times post-infection. Using confocal microscopy, we generated z-stacks to visualize the inside of the ovarioles and testes. Indeed, Y. pestis was readily observable, appearing to localize inside the ovarioles rather than on the outside surface (Fig. 2A-D). Similarly, we readily observed tomato-fluorescing Y. pestis in the testes of male adult fleas (Fig. 2E-H). Therefore, perhaps not unexpectedly, Y. pestis was also found in the spermatheca (Fig. 2I-L). Other tissues that contained Y. pestis that was visible by confocal microscopy included Malpighian tubules, but not salivary glands (Extended Data Fig. 1, salivary glands not shown). Furthermore, Y. pestis could be seen in the reproductive organs up to 15 weeks after the adult fleas were infected (Extended Data Fig. 2). This result suggests that Y. pestis infection of the reproductive tissue is stable throughout the lifespan of the adult flea.

Viable Y. pestis is present in the oviposited eggs and larvae derived from infected adult fleas.

To determine whether viable Y. pestis was present in the eggs, we devised an egg collection chamber, using sifting to separate the eggs from bedding debris. Adult fleas that are feeding regularly have a steady reproduction at 3.6 to 5.6 eggs per female per day which decreases as the fleas age ^26,27. Eggs from X. cheopis are 477–504 µm long with a width of 297–333 µm and off-white in color. Sifting was carried out every 1–3 days and eggs were collected of the expected size and color. Eggs were thoroughly washed and then processed for imaging or for plating to determine bacterial titer. Control fleas, given blood that was not infected, produced eggs with no fluorescent signal either on or within the egg tissue (Fig. 3A-D). In sharp contrast, tdTomato fluorescence was abundant in the interior of eggs produced by Y. pestis-infected fleas, localizing to nearly the length of the egg (Fig. 3E-L). We found prominent fluorescent signal in more than half of the eggs that were tested, whether they were collected 3 (Fig. 3D-F) or 7 (Fig. 3I-L) days post-infection. Considering blood intake initiates copulation and subsequent oviposition, the data suggest that egg-derived Y. pestis were either transferred directly to the egg due to midgut escape during the bloodmeal, or that colonization of the spermatheca results in the continuous opportunity for Y. pestis to infect newly developing eggs ²⁸.

We also collected larvae by sifting and found 5 of 8 were positive, with tdTomato fluorescence that appeared to be present in nearly the entire length of the larvae (Fig. 4D-I) compared to larvae from control fleas which were uninfected (Fig. 4A-C). To rule out the likelihood that larvae were infected by eating Y. pestis that had been expelled from the hindgut, we collected eggs that were deposited by Y. pestis-infected adult X. cheopis and reared them in a separate sterile container into larvae. As expected, fluorescent Y. pestis was readily visualized in these samples (Fig. 4G-I). Together, these data indicate that Y. pestis in the eggs survives development into the larval stage. Additional eggs from Y. pestis-infected adult fleas were collected, placed into sterile sawdust with sterile larvae food where they developed into pupae and F1 adults. Pupae exhibited a high degree of autofluorescence (Extended Data Fig. 3A-B). Nevertheless, strong tdTomato expression could be detected in the pupae reared from Y. pestis-infected eggs (Extended Data Fig. 3C-D). Likewise, fluorescent Y. pestis could also be visualized in the midguts of the F1 adult fleas (Extended Data Fig. 4A-D). Bacteria harvested from these fleas were screened for Y. pestis plasmid-encoded genes, caf1 and pla by PCR, and indeed, the transovarially-transmitted bacteria were positive for both genes, suggesting they retained the extrachromosomal plasmids (Extended Data Fig. 4E).

Transovarially acquired Y. pestis is competent for transmission.

To determine whether transovarially transmitted bacteria were viable, we compared bacterial titers in eggs, larvae, pupae and F1 progeny. The median number of colonies recovered from eggs was only 5 CFU (Fig. 5A). In the later stages of development, bacteria appeared to replicate, with median titer recovered in larvae of approximately 50 CFU, and in pupae, the median titer was over 100 CFU. In the newly emerged adult F1 fleas collected prior to their first blood feeding, two midguts were harvested and combined for plating, resulting in 173 CFU recovered. To confirm the retention of extrachromosomal plasmids, bacterial stocks were made and then used for PCR to amplify the Y. pestis plasmid-specific genes caf1 and pla. Following PCR, the expected-size bands were purified and processed to determine the DNA sequence in order to conclusively confirm Y. pestis (Extended Data Fig. 4E, sequence data not shown). Together, these data indicate that transovarially transmitted Y. pestis remain viable throughout the developmental cycle and the recovery of viable Y. pestis from the midgut of F1 adult fleas suggests that transovarially transmitted bacteria are competent for transmission.

We therefore evaluated whether Y. pestis isolated from eggs remained competent for flea transmission to a mammalian host. Naïve adult fleas were challenged with egg-isolated Y. pestis in the artificial membrane feeder and on days 3 and 7, infected fleas were used in a transmission study. On day 3 post-infection, it was evident that egg-isolated Y. pestis had been successfully transmitted, and its localization to midgut biomass appeared indistinguishable from fleas infected with the original Y. pestis stock (Fig. 5B-C). Similar kinetics of egg-isolated Y. pestis were observed in the flea midgut as seen for the parent Y. pestis strain with similar bacterial load observed on day 7 post-infection. Overall, these data suggest that transovarially transmitted Y. pestis is competent for re-entry into the sylvatic plague cycle.

Transmission electron microscopy (TEM) shows Y. pestis localized to reproductive tissue in adult fleas.

To confirm that bacteria were present in the testes and ovaries of infected adult fleas, we used transmission electron microscopy to image tissues that were dissected from male and female fleas. In the females, we observed bacteria in the ovaries, and even in the safety pin morphology that is characteristic of Y. pestis (Fig. 6B-C). These bacteria were not found in fleas that were not infected (Fig. 6A). In the male fleas, we readily identified abundant Y. pestis in the basal membrane of the testes, appearing on the distal side of the spermatocyte follicle, often within membrane-enclosed vesicles (Fig. 6D-F). This suggests that Y. pestis may be intracellular in the reproductive tissue. Overall, the TEM confirms that Y. pestis exit the midgut and localize to the reproductive organs of an adult flea.

In this work, we have provided multiple layers of evidence that transovarial transmission occurs in X. cheopis infected with Y. pestis. These findings provide visual, quantitative, and genetic support that low levels of Y. pestis escape from the midgut throughout the lifespan of an infected flea, entering the reproductive tract where they are able to colonize progeny eggs. In the more than 100 years of Y. pestis research, escape from the alimentary tract has never been reported. Previous studies were heavily focused on salivary glands, and in agreement with these results, we did not observe any Y. pestis in the salivary glands. Furthermore, the amount of colony forming units harvested from flea eggs is exceedingly small, averaging fewer than 10 CFU, consequently it may simply be that previous experiments were not sensitive enough to capture the low level of transovarial transmission. In this work, the high-level expression of the tdTomato reporter revealed a new understanding of sylvatic plague.

Fleas are vectors of other diseases, namely rickettsiosis and cat scratch fever caused by Rickettsia felis or typhi or Bartonella henselae, respectively. For Rickettsia typhi, transovarial transmission was observed in fleas more than 30 years ago ²⁹. Like Rickettsia felis, intracellular Y. pestis appear to localize to the ovariole tissue ³⁰. However, there are likely significant differences in the underlying mechanism of midgut escape. Whereas R. felis infection of the midgut epithelium was readily apparent shortly after infection, there was no detectable infection of these cells by Y. pestis. Following invasion of the midgut epithelium R. felis was observed in hemocoel prior to its systemic dissemination to the reproductive tract and salivary glands. In contrast, Y. pestis was not observed in hemocoel or salivary glands.

Transovarial transmission could be a contributor to maintenance of Y. pestis within the environment. From the data shown here, viable Y. pestis is continuously deposited in eggs, suggesting most, if not all, progeny fleas remain infected. Furthermore, the strong selective pressure of the sylvatic plague cycle would favor the retention of virulence factors by transovarially-transmitted Y. pestis. Therefore, it seems likely that transovarial transmission could sustain low levels of Y. pestis in a rodent community. Environmental factors such as precipitation and temperature are predicted to have an impact on the transovarial transmission cycle, thereby having multi-layered contributions to the prevalence of Y. pestis in the environment. By understanding transovarial transmission, we may improve our ability to model the enzootic cycle of plague.

Acknowledgements: We wish to give special thanks to Dr. Alexander Jurkevich for expert technical guidance with the confocal imaging and to DeAna Grant for conducting the transmission electron microscopy processing and imaging. We are also grateful to Dr. Paul Anderson and Jenna Canfield for helpful discussion and critical comments on the manuscript. Sequencing was performed by the University of Missouri Genomics Technology Core; microscopy was conducted at the University of Missouri Advanced Light Microscopy Core; and transmission electron microscopy was conducted at the Electron Microscopy Core at the University of Missouri. This work was partially supported by PHS NIH/NIAID 1R21AI178547 and the University of Missouri System Tier 2 Strategic Investment Program.

Author contributions: C.P. designed and performed experiments, interpreted data and co-wrote the manuscript; B.B. and Q.S. contributed to data interpretation and editing of the manuscript; D.A. contributed to experimental design and interpretation, and co-wrote the manuscript. All authors approved the manuscript.

Competing interests: The authors declare no competing interests.

Additional information: Supplementary information is available for this paper.

Data Availability:

The sequence chromatographs and alignments, image data sets and corresponding metadata that support the findings of this study are available in Figshare with the identifier 10.6084/m9.figshare.c.6845496. [37]

All unique biological materials, including bacterial strains and plasmids, are available from the corresponding author upon request. There are no restrictions on any of these materials.

Materials and Correspondence: Correspondence and requests for materials should be addressed to Dr. Deborah Anderson, [email protected].

Reprints and permissions information is available at www.nature.com/reprints.

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Animals: Neonatal and adult mice were used in these experiments for colony maintenance blood feeding as well as preparation of the artificial feeder, according to previously published protocols ^31,32. Colony fleas were provided opportunity for blood feeding 2-3 times/week prior. The blood feeding and infection protocol were approved by the University of Missouri Animal Care and Use Committee.

Infection: These experiments were conducted with a laboratory strain of Yersinia pestis KIM6+, which lacks the type III secretion system plasmid pCD1 ³³. The recombinant derivatives of KIM6+ used in this work did not reintroduce the pCD1 plasmid, and therefore are classified as select agent-exempt strains by the US Center for Disease Control and Prevention. Prior to use in infection, laboratory reared, naive X. cheopis were separated from the colony and starved for at least five days and no more than seven days to improve feeding efficiency during infection. Groups of 50 fleas were infected with Y. pestis strain KIM6+ carrying plasmid-expression of the fluorescent protein tdTomato (Excitation: 554, Emission 581) ³⁴. An artificial membrane feeder was constructed using skin from an adult mouse. Blood was inoculated with 5x10⁸ to 1x10⁹ Y. pestis and maintained at 37°C, and fleas were allowed to feed for 1 hour. The species of animal blood (rat, mouse, pig, or prairie dog) used in the artificial feeder is indicated in the figure legends. When fleas were removed from the feeder, they were observed to determine intake of the bloodmeal. Fleas that had not fed were removed from the study.

Midgut processing: Fleas were euthanized on days 1, 3 and 7 post-infection without additional blood feeding. For experiments that lasted more than 7 days, fleas were provided maintenance blood meal every 7 days throughout the duration. Dissected midguts and other reproductive tissues were isolated and placed onto sterile slides, fixed with 4% paraformaldehyde, and rinsed 3 times with PBS for a minimum of 30 minutes each time prior to mounting.

Confocal microscopy: Midguts, eggs, larvae and pupae were imaged using a Leica SP8 confocal microscope. For quantification of midgut biomass, images were converted to 8-bit grayscale for quantification of observed integrated density (ID_O). Control fleas (n=10), fed in parallel with uninfected blood and analyzed 1 day after feeding, were used to determined background fluorescence. Background signal and midgut area were used to normalize the samples and calculate integrated density (ID= ID_O - (midgut area x background)). Image J software was used to capture images and quantify the signal intensity, reported as relative fluorescent units (RFU) ³⁵.

Quantification of bacterial load: For plating, individual samples were homogenized in 10μL sterile PBS, midguts were pooled in groups of 3; serial dilutions were performed in sterile PBS and all dilutions were plated in duplicate onto heart infusion agar (HIA) or Yersinia selective agar (YSA). Isolated colonies from eggs, larvae, pupae, and F1 adults were streaked for isolation before storing in bacteriological freezing media at -80°C.

Egg collection: Infected fleas using rat blood were maintained in modified housing with 300-micron mesh on the lid, 550-micron mesh under the bedding, and a lower chamber that could be easily removed. Eggs and larvae were collected by sifting, with care to avoid contact with infected feces and fleas. Fleas were maintained in the modified container housing and provided uninfected rat bloodmeal every 7 days. Sifting for eggs and larvae occurred every 1-3 days post-infection. To aseptically collect the eggs and larvae, a sterilized cotton applicator was moistened with double-distilled, sterile H₂O and used to pick up eggs or larvae. These specimens were placed onto a sterile slide, washed in sterile PBS three times, observed between washing to ensure no materials or feces were in contact, then mounted in 35% glycerol for confocal microscopy or transferred to sterile PBS. After washing three times in sterile PBS, a new sterile, moistened cotton applicator was used to transfer the egg to an agar plate. Each egg was punctured with a sterile needle, releasing the contents onto the agar. Similarly, larvae were washed three times with sterile PBS, then homogenized and plated.

Development of Y. pestis-infected eggs or larvae: Eggs were isolated, washed in sterile PBS, and placed into a sterile flea chamber with mesh on the top and bottom containing sterile sawdust and larval food. These samples were imaged or plated on agar after development into larvae, pupae, or F1 adults.

PCR and genome annotation: Bacteria that were isolated from eggs were grown overnight and DNA was isolated using a Quick-DNA® Microprep kit (Zymo Research, California, USA). Conventional PCR amplification of caf1 and pla were performed using the primers shown in Extended Data Table 1. All positive PCRs were confirmed by sequencing; sequenced nucleotides were aligned in Geneious Prime using MAFFT ³⁶.

Transmission study: Transmission assays were carried out as previously described for fleas infected with KIM6+ptdTomato ⁸. For assays using bacteria originally harvested from eggs, minor modifications were made. Briefly, egg isolated bacteria were used to infect adult fleas on an artificial feeder. On day 3 or 7 post-infection, groups of 10 fleas were used for the transmission assay and fed on uninfected rat bloodmeal in the artificial membrane feeder for 1 hour. Blood and skin were processed to quantify Y. pestis by plating to determine the number of bacteria transmitted per group.

Transmission electron microscopy. Following the same methods, fleas were either fed an uninfected rat bloodmeal or a Y. pestis-infected rat bloodmeal and then groups of 10 were separated based on sex. On day 3, fleas were euthanized, and the ovaries, testes, and midguts were dissected, fixed in 2% paraformaldehyde, 2% glutaraldehyde in 100 mM sodium cacodylate buffer with a pH of 7.35. Each sample was allowed to settle, and the resulting tissue was resuspended in Histogel (ThermoScientific, Kalamazoo, MI). Tissues were rinsed in 100 mM sodium cacodylate buffer with a pH of 7.35 containing 130 mM sucrose. Secondary fixation was performed using 1% osmium tetroxide (Ted Pella, Inc. Redding, California) in cacodylate buffer. Specimens were incubated at 4°C for 1 hour, then rinsed with cacodylate buffer and further with distilled water. En bloc staining was performed using 1% aqueous uranyl acetate and incubated at 4°C overnight, then rinsed with distilled water. A graded dehydration series was performed using ethanol, transitioned into acetone, and dehydrated tissues were then infiltrated with Epon resin and polymerized at 60°C overnight. Sections were cut to a thickness of 75 nm using an ultramicrotome (Ultracut UCT, Leica Microsystems, Germany) and a diamond knife (Diatome, Hatfield PA). Images were acquired with a JEOL JEM 1400 transmission electron microscope (JEOL, Peabody, MA) at 80 kV on a Gatan Rio CMOS camera (Gatan, Inc, Pleasanton, CA). All samples were prepared, processed and imaged at the University of Missouri Electron Microscopy core.

Statistical analysis. Data was grouped based on trial and microscopic analysis of individual flea midguts and tissue, with controls groups derived from the parent Y. pestis and uninfected fleas. Descriptive statistics were evaluated using SPSS 26 and graphed using OriginPro.

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Transovarial transmission of Yersinia pestis in its flea vector, Xenopsylla cheopis

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