Abstract
The γ-proteobacterium Francisella tularensis causes seasonal tick-transmitted tularemia outbreaks in natural rabbit hosts and incidental infections in humans in the south-central United States. Although Dermacentor variabilis is considered a primary vector for F. tularensis, Amblyomma americanum is the most abundant tick species in this endemic region. A systematic study of F. tularensis colonization of A. americanum was undertaken to better understand its potential to serve as an overwintering reservoir for F. tularensis and as a bridging vector for human infections. Colony-reared A. americanum were artificially fed F. tularensis subspecies holarctica strain LVS via glass capillaries and colonization levels determined. Capillary-fed larva and nymph were initially infected with 104 CFU/tick which declined prior to molting for both stages, but rebounded post-molting in nymphs and persisted in 53% at 103 to 108 CFU/nymph at 168 days post-capillary feeding (longest sampling time in the study). In contrast, only 18% of adults molted from colonized nymphs maintained LVS colonization at 101 to 105 CFU/adult at 168 days post-capillary feeding (longest sampling time). For adults, LVS initially colonized the gut and disseminated to salivary glands by 24 h and had an ID50 of <5CFU in mice. Francisella tularensis infected the ovaries of gravid females, but transmission to eggs was infrequent and transovarial transmission to hatched larvae was not observed. The prolonged persistence of F. tularensis in A. americanum nymphs supports A. americanum as an overwintering reservoir for F. tularensis from which seasonal epizootics may originate; however, although the rapid dissemination of F. tularensis from gut to salivary glands in adults A. americanum is compatible with intermittent feeding adult males acting as bridging vectors for incidental F. tularensis infections of humans, acquisition of F. tularensis by adults may be unlikely based on adult feeding preference for larger mammals which are not involved in maintenance of sylvatic tularemia.
Introduction
The γ-proteobacterium Francisella tularensis subspecies tularensis can cause severe systemic infections in natural rabbit hosts and incidental infections in humans in its endemic region in the south-central United States [1–3]. In this endemic region, tularemia is primarily a tick-vectored disease with a spring-summer seasonal pattern corresponding to the questing period of its tick vectors [1]. The American dog tick Dermacentor variabilis is thought to be the primary vector for F. tularensis in its natural rabbit host in the endemic region as well as a possible overwintering reservoir for F. tularensis, although the lone star tick Amblyomma americanum is the most abundant tick species in this endemic region. Infestation of D. variabilis on small to medium-sized wild mammals has been associated with sylvatic tularemia, and D. variabilis has been demonstrated to be an experimental vector for F. tularensis and to support transstadial transmission [1,4–7]. However, the host feeding preference of A. americanum for larger mammalian hosts including humans may make this tick species more competent vector for transmission of the bacterium to humans [1,8].
Amblyomma americanum has been speculated to transmit F. tularensis in nature and is a competent experimental vector exhibiting transstadial transmission [1,9]. Although F. tularensis has been detected in wild-caught A. americanum, its role in transmission of tularemia in nature has not been firmly established [9–11]. To better understand the potential role of A. americanum in transmission of tularemia, a detailed systematic study of quanta and duration of F. tularensis colonization of larvae, nymph and adult A. americanum was undertaken. In addition, the distribution of F. tularensis in tick tissues of nymph and adults was determined as well as examination of transovarial transmission and transmission to rodents via tick saliva. These studies were accomplished using an artificial colonization method of feeding F. tularensis to various tick stages using glass capillary tubes placed over the tick mouthparts.
We had previously used this method to study the biology of F. tularensis in D. variabilis. In that study, we found that F. tularensis can infect D. variabilis as larvae or nymphs and maintain colonization through the adult stage, and infected D. variabilis can transmit infection to susceptible hosts, making D. variabilis a potential overwintering reservoir for F. tularensis [5]. However, the relative high abundance of A. americanum compared to D. variabilis in the tularemia endemic region raises the question of what role A. americanum may play as a vector of F. tularensis in transmission and persistence of the agent in its endemic region.
Although D. variabilis and A. americanum are both ixodid three-host ticks, they differ in many aspects of their lifestyles and natural history which may impact their potential for maintenance and transmission of F. tularensis in the tularemia endemic region. Dermacentor variabilis larvae and nymphs preferentially feed on rodents and medium-sized mammals from which it can acquire F. tularensis, whereas only the larval stage of A. americanum feed on rodents or medium-sized mammals from which it might acquire F. tularensis [12]. These ixodid ticks typically have two to three-year lifespans with D. variabilis larvae and adults and A. americanum nymphs surviving the winter in the tularemia endemic region [12–14]. These differences in lifestyles and abundance may result in different roles for these tick species in maintaining and transmitting F. tularensis in its endemic region, particularly if the biology of F. tularensis also differs in these ticks.
Although previous studies of F. tularensis in A. americanum provided data on infection quanta and demonstrated transstadial transmission [10], our study provides additional detailed systematic studies regarding duration and quanta of colonization as well as examining the tissue localization and transovarial transmission of F. tularensis in A. americanum, the latter of which have not previously been reported. For the current study, artificial glass capillary tube feeding method of colonizing ticks with Francisella was used to colonize A. americanum with F. tularensis subspecies holarctica strain LVS. We found that A. americanum nymphs which acquired F. tularensis colonization as larvae are able to maintain colonization for sufficient duration and at sufficient quanta to potentially maintain F. tularensis in the endemic region from one season to the next; however, the feeding preference of A. americanum nymphs and adults for larger mammals may reduce its importance for maintenance of the tularemia enzootic cycle in rabbits but may facilitate its role as a bridging vector for tularemia in humans.
Materials and Methods
Ticks, bacterial strains and growth conditions
A. americanum larvae, nymphs, and adults were obtained from the Tick Rearing Facility, Department of Entomology and Plant Pathology, Oklahoma State University (Stillwater, OK). Larvae were collected after they were fed to repletion on rabbits. Nymphs were partially fed on sheep until they weighed ≥4.5 mg/nymph. Unfed flat adult ticks were used for quanta and duration of LVS colonization studies and for tissue distribution and infectivity studies. Adult ticks used for salivary induction experiments were partially fed on sheep for five to six days. Female ticks used to study transovarial transmission were fed to repletion on sheep. The tick-feeding on animals were carried out in strict accordance with the recommendations of Institutional Animal Care and Use Committee, Oklahoma State University. The protocol was approved by the Committee (IACUC protocol AG-50-219).
Francisella tularensis subsp. holarctica strain LVS was supplied by the Oklahoma State Department of Health. A green fluorescent protein (GFP) expressing transformant of LVS was prepared by electroporation of a GFP plasmid (pFNLTP6 gro-gfp), gift of Thomas C. Zhart (Medical College Wisconsin, Milwaukee, WI), into LVS [15]. Preparation of inocula and growth of LVS has been described earlier [5]. Francisella tularensis was grown on chocolate agar plates (Hardy Diagnostics, Santa Monica, CA.) at 37°C in 5% CO2 for 72 h. BBL Prompt Inoculation System (BD Diagnostics, Franklin Lakes, NJ) was used to spike F. tularensis in the tick inoculum. All chemicals used in the study were purchased from Sigma (St Louis, MO) unless indicated otherwise.
Capillary tube feeding of larvae, nymphs and adult ticks
The surface disinfection of ticks and the capillary tube-feeding technique have been described earlier [5]. For feeding each batch of ticks, fresh medium was made and spiked with a uniform inoculation dose of 107 CFU/ml LVS. The larvae, nymphs and adults were fed for 18 hours at 30°C and 90% relative humidity. Capillary tube feeding of nymphs and adult ticks was initially assessed by weighing ticks before and after feeding. Only those ticks (nymphs and flat adults) which gained ≥0.3 mg after capillary tube feeding were used for the experiment unless otherwise specified. Based on the estimated minimum tick meal ingested for a ≥0.3 mg body weight gain and the inoculum dose of 107 CFU LVS/ml, the minimum post-capillary feeding infection level is calculated to be 3 × 103 CFU/tick. After feeding, ticks were either surface disinfected by washing for 5 seconds each in 30% hydrogen peroxide, distilled water and 70% isopropyl alcohol and minced for determination of CFUs or were maintained in microcentrifuge tubes capped with moistened cotton plugs and kept in a humidity chamber (relative humidity of > 90%) at 22±1°C (unless specified otherwise) with automated artificial lighting to simulate a 12 h day-night cycle. Determination of CFU per tick was done by sacrificing groups of ticks which were analyzed individually at one day post-capillary feeding and then at an interval of 7 days up to the longest sampling time in these studies of 168 days. To determine the bacterial numbers in ticks at various times of colonization, individual ticks in microcentrifuge tubes were minced with a sterile sharp pointed tweezers, bacteria extracted by incubation in PBS containing 64 μg/mL ampicillin for 2 h at room temperature on a rotor platform mixer (Boekel Scientific, Feasterville, PA.), serially diluted in PBS containing 64 μg/mL ampicillin, and plated on chocolate agar plates for CFU determination following incubation at 37°C in 5% CO2 for 72 h. To determine the bacterial numbers in tissues at varying periods of post-capillary tube feeding, individual ticks were dissected into gut, salivary glands, and ovaries under sterile conditions using a dissecting microscope. The tick tissues were placed in PBS containing 64 μg/mL ampicillin and processed as described above for whole ticks. Hemolymph from ticks was collected into sterile glass capillary tubes from the cut ends of the tick’s legs.
Immunohistochemistry
For immunohistochemical analysis, nymphs which were held in humidity chambers for 40 or 168 days post-capillary tube feeding were fixed in Carson’s fixative, embedded with paraffin and sectioned and affixed to glass slides. After deparaffinizing, the sections were incubated with phosphate buffered saline with 0.05% Tween 20 (PBST) at room temperature for 15 min. and then incubated at 37°C for 1 h with F. tularensis antiserum (Beckton Dickinson, Sparks, Maryland) at 1/60 dilution in PBST. Antiserum preabsorbed against LVS and uninfected tick sections were used as negative controls (S1 Fig). After washing the slides with PBST five times followed by a final washing with distilled water, the sections were incubated with fluorescein isothiocyanate (FITC) conjugated secondary antibody (KPL, Gaithersburg, MD) at 1/60 dilution in PBST at 37°C for 30 min. The sections were then washed in PBST twice, PBS once and finally washed with distilled water. The slides were visualized using Nikon Eclipse 50i epi-fluorescence microscope and Nikon digital sight DS-5M-L1 digital camera. For visualizing LVS in tick hemolymph, adult ticks were capillary tube fed with LVS expressing GFP. At 1 and 4 weeks post-capillary tube feeding, hemolymph was collected, placed directly on glass slide with coverslip, and visualized using the epi-fluorescent microscope.
Real-time quantitative PCR
Real- time quantitative PCR was conducted to confirm the absence of LVS in larval and adult ticks and egg masses in which LVS was not detected by microbial culture. Each sample was analyzed using Fast SYBR green master mix on an AB 7500 Fast Real-Time PCR System (Applied Biosystems, Foster City, CA.). A negative control (no template) was processed for each analysis. Samples were assessed to be positive when a 97bp band F. tularensis ISFtu2 amplification product [16] was confirmed by sequencing and analyzing the dissociation curve. The primer sequence used for the RT-PCR reaction did not detect the presence of F. tularensis or Francisella-like endosymbionts in the colony reared larvae, nymphs or adult ticks prior to feeding ticks LVS. The RT-PCR reaction (20 μL) contained 10 μL Fast SYBR green master mix, 6 μl DNase RNase-free water, 1 μL forward primer (ISFtu2F), 1 μL reverse primer (ISFtu2R) and 2 μL template. Cycling conditions were 95°C for 20 seconds, followed by 34 cycles of 95°C for 10 seconds and 60°C for 30 seconds. Total DNA from each tick (tick minceate in 100 μl PBS) was extracted using DNeasy Tissue Kit (Qiagen, Valencia, CA.), with a final elution volume of 50 μL.
Infection of ticks by intra-hemocoelic (i.h.) inoculation
To determine the lowest infectious inoculation dose for adult ticks resulting in detection of LVS and to infect gravid females and partially fed adult ticks, 1 μL of the inoculum containing LVS in PBS was injected i.h. in the ventral region of the tick, medial to the caudal most coxa using 10 μL custom-made Hamilton syringe with a 0.5 inch, 33 gauge needle (Hamilton Company, Reno, NV).
Detection of LVS in tick saliva
To detect LVS in salivary secretion of ticks infected with LVS via capillary tube-feeding, these ticks were induced to secrete saliva by injection with a secretagogue. Two days post-capillary tube feeding, ticks were immobilized dorsal side on the sticky part of duct tape placed on a double-sided adhesive tape. Ticks were then injected i.h. with 4 μL of 1 mM dopamine, 1 mM theophilline and 3% DMSO in PBS (pH 7.3) [17] every 15 min for 1 h. Saliva was collected in a 10 μL (internal diameter of 0.0219 inch) volume glass capillary tubes (Drummond Scientific Company, Broomall, PA) placed over the hypostome of the tick. The capillary tube for collecting saliva was held in place using modeling clay.
Infective dose 50 in BALB/c mice
The Animal Care and Use Protocol for mouse ID50 experiments was approved by the Oklahoma State University Institutional Animal Care and Use Committee (IACUC protocol VM-10-1). To determine infectivity of LVS recovered from tick salivary secretions, eight partially fed adult ticks (colonized with LVS four days previously via capillary tube feeding and held at 27°C) were induced to secrete saliva under sterile conditions, and the saliva from all eight ticks were pooled in 200 μL PBS containing 64 μg/mL ampicillin. The saliva in PBS-ampicillin was diluted to make particular inoculum doses. Five experimental groups of BALB/c mice (six mice in each group) were injected i.p. with 5 CFU, 50 CFU, 500 CFU, 5x103 CFU, and 1x104 CFU LVS from the pooled and diluted saliva. Another five experimental groups of BALB/c mice (six mice in each group) were injected i.p. with 0.4 CFU, 4 CFU, 39 CFU, 194 CFU and 387 CFU of laboratory cultured LVS. One control group of six mice were injected with pooled and diluted tick saliva in PBS-ampicillin from uninfected ticks (capillary tube fed with tick meal without LVS four days previously and held at 27°C). All the mice that showed the clinical endpoint (ruffled haircoat, huddling, lethargy, and decreased mobility) were euthanized. Liver and spleen were aseptically removed form the mice, weighed and homogenized. Blood was collected from the heart immediately after euthanizing the mice, serial 10-fold dilutions made and plated on chocolate agar plates, and CFUs were determined after 72 h of incubation at 37°C and 5% CO2. The data from the experiment were used to calculate ID 50 using Reed-Muench method [18].
Transovarial transmission of LVS in infected gravid adult female A. americanum to larvae via the egg stage
To assess whether female A. americanum transovarially transmit F. tularensis to hatched larvae, adult females fed on sheep to repletion were inoculated via i.h route one day post-repletion either with 1μL of 107 CFU/mL LVS in PBS or with 1μL PBS alone (control). Eggs oviposited between days 10 to 50 post injection were collected and weighed. These eggs after surface sterilization (washing for 5 seconds each in PBS and 70% isopropyl alcohol) were minced and the lysate were serially diluted in PBS containing 64 μg/mL ampicillin and plated for CFU determination. At the completion of oviposition adult female ticks were surface sterilized and hemolymph was collected. The ticks were dissected and tissues including gut, Malpighian tubules, and ovaries were separated out minced and the lysate were serially diluted in PBS-ampicillin and plated for CFU determination. Subsets of the oviposited eggs were placed in microcentrifuge tubes capped with moistened cotton plugs and kept in a humidity chamber for hatching. The hatched larvae after surface sterilization were minced and the lysate were serially diluted in PBS-ampicillin and plated for CFU determination.
Statistical analysis
Francisella tularensis infection levels in different groups of A. americanum ticks during adult colonization, transstadial transmission from larva to nymphs, and nymph to adult were compared by using 1-way analysis of variance on log-transformed data followed by pairwise multiple comparison of mean CFU values using Holm-Sidak tests. Overall significance level for Holm-Sidak tests was P = 0.05. Student's t-test was performed to determine the statistical difference in the mean CFU/infected tick between molted adult male and female A. americanum. All statistical analyses were performed with SigmaPlot v11.0 Graphics software (Systat Software Inc., Chicago, IL).
Results
LVS infection of A. americanum larvae and transstadial transmission to nymphs
To assess quanta and duration of infection in larval A. americanum and transstadial transmission of F. tularensis to nymphs molted from infected larvae, larvae fed to repletion on rabbits were exposed to LVS by capillary feeding tick meal containing 107 CFU/mL LVS. On day one and every seventh day through day 168 post-capillary feeding, groups of larvae or nymphs molted from larvae were assayed individually for LVS (Fig 1). The degree to which larvae took in the capillary fed meal was not ascertainable by weight change in larvae, but at one-day post-capillary tube feeding, LVS was detected with a mean infection level of 1.5 × 104 CFU/larva in 100% of larvae (Fig 2; limit of detection was 2 CFU/larva). For our experimental holding conditions (relative humidity = 95% and mean room temperature of 22±1°C), the larvae molted to nymphs between day 21 and 28 post-capillary tube feeding. Around the time of molting, both the % infected and the mean bacterial count declined (9.6±4.8 × 101CFU/tick, Fig 2). At molting, LVS was detected using immunohistochemistry in gut and in salivary tissue (Fig 3A and 3B). The % infected nymphs remained between 50 and 90% post-molt, but quanta of infection increased to 102 to 103 CFU/nymph between molting and 42 days post-molting, and then increased in level to reach ≈1 × 105 CFU/nymph at 70 days post-capillary feeding (49 days post-molting). This mean level of colonization was maintained through the longest sampling time of 168 days (147 days post-molting) (Fig 2A). Significant difference in the mean colonization levels between molting (day 21) and 70, 77, 84, and 168 days post-capillary feeding (unadjusted P < 0.001) was observed. There were no significant differences between any other time points. At 147 days post-molting, LVS was detected only in gut tissue (Fig 3C and 3D). Similar overall survival percentages of 63% and 65% were observed for larvae and nymphs molted from larvae fed LVS compared to those fed tick meal alone.
Fig 1. Quantum and duration of colonization and transstadial transmission of LVS experimental design.
Larvae, nymphs and adults A. americanum in batches of various sizes were capillary fed tick meal containing LVS. Ingestion of capillary fed tick meal was determined for nymph and adult ticks by post-feeding weight gains of ≥0.3mg. Nymphs and adults showing <0.3mg weight gains were excluded from further analysis. The overall rate of ticks positive for LVS for each tick life cycle stage capillary tube fed LVS is given as total number of ticks which were positive for LVS (+LVS)/total number of ticks tested.
Fig 2. LVS transmitted transstadially from larvae to nymphs.
Open circles are capillary tube fed larvae, and filled circles are molted nymphs. The calculated mean CFU/tick for colonized larvae and molted nymphs for each time point is represented by the horizontal line. For each time point, n was 15. (B) Percentage of colonized ticks in the same experiment.
Fig 3. Visualization of LVS in colonized nymphs at 40 and 168 days post-capillary tube feeding.
(A) Wright Giemsa stained section of an LVS colonized nymph which was capillary tube fed LVS as a larva, molted to nymph, and was held at 22±1°C for 40 days. (B) Immunostained section of the same nymph. LVS colonization of both the salivary glands and gut was seen. (C) Wright Giemsa stained sections of an LVS colonized nymph which was capillary tube fed LVS as a larva, molted to a nymph, and was held at 22±1°C for 168 days. (D) Immunostained section of the same nymph. LVS colonization was observed only in the gut. Numbers identify (1) muscle, (2) salivary gland, (3) gut, (4) exoskeleton and (5) Malpighian tubule. 200x magnification.
LVS infection of A. americanum nymphs and transstadial transmission to adults
To assess quanta and duration of infection A. americanum nymphs and transstadial transmission of F. tularensis in A. americanum adults molted from infected nymphs, nymphs fed for 4 days on sheep were exposed to LVS by capillary feeding tick meal containing 107 CFU/mL LVS. The degree to which nymphs ingested tick meal was ascertained by comparison of pre-feeding and post-feeding weights in which 68% of fed nymph gained ≥0.3 mg/nymph weight gain post-feeding (Fig 1). Sampling only those nymphs with ≥0.3 mg/nymph weight gain post-feeding, on day one LVS was detected in 100% of nymphs with a mean level of 1.3±0.01 × 104 CFU/nymph (Fig 4A). The percent nymphs infected declined towards molting to adults at 35 days, at which time 8/15 adults molted from infected nymphs were positive for F. tularensis by microbial culture, demonstrating the transstadial transmission of F. tularensis in capillary tube fed A. americanum (Fig 4A). There was a progressive decline in the percent of molted adult ticks infected by F. tularensis to 13% by post-capillary feeding day 56 (day 21 post-molting) and remained at this % at longer sampling times (Fig 4B). Like nymphs molted from infected larvae, the adults molted from infected nymphs showed a decline in quanta of infection until molting, but then the quanta of infection trended higher post-molting (Fig 4A). However there were no significant statistical differences between any time points. The overall survival rate for nymphs and adults molted from nymphs which had increased ≥0.3 mg/nymph weight gain post-feeding was 88% (Fig 1).
Fig 4. LVS transmitted transstadially from nymphs to adults.
(A) Open circles are capillary tube fed nymphs, and filled circles are molted adults. The calculated mean CFU/tick for colonized nymphs and molted adults for each time point is represented by the horizontal line. For each time point up to day 168, n was 15. (B) Percentage of colonized ticks in the same experiment.
LVS infection of A. americanum adults
To assess quanta and duration of infection of F. tularensis in A. americanum adults, flat adults which had not been fed a blood meal as adults were exposed to LVS by capillary feeding tick meal containing 107 CFU/mL LVS. The degree to which adults ingested tick meal was ascertained by comparison of pre-feeding and post-feeding weights in which 62% of fed adults gained ≥0.3 mg/adult weight gain post-feeding (Fig 1). On days one through 28 (Fig 5) and on day 168 post-capillary feeding, groups of adults were assayed for LVS. Sampling only those adults with ≥0.3 mg/tick weight gain post-feeding, a range of mean LVS infection quanta detected were 102 to 105 CFU/tick for the 28 days post-capillary tube feeding period. For the longest sampling time of 168 days post-capillary feeding, 7/29 of flat adults which gained ≥0.3 mg/tick weight gain post-feeding maintained infection with quanta of infection ranging from 101 to 106 CFU/tick. However there were no significant statistical differences between any time points. The overall survival rate for adults which had increased ≥0.3 mg/nymph weight gain post-feeding was 57% (Fig 1).
Fig 5. LVS colonization of adult A. americanum.
The filled circles are capillary tube fed adults. The calculated mean CFU/tick for colonized unfed adults for each time point is represented by the horizontal line. For each time point, n was 11; for day 168, n was 29.
Determination of inoculum dose of LVS necessary to establish infection of A. americanum adults
To determine a minimum infectious dose of F. tularensis necessary to cause infection of adults ticks, flat adults were injected intra-hemocoelically (i.h.) with 140, 13 or 2 CFUs of LVS. At the high inoculum dose, 10/10 ticks were positive for LVS on day 7 post-injection with a mean quanta of 2.5 × 103 CFU/tick; at a dose of 13 CFU/tick, 8/10 tick were positive on day 7 with a mean quanta of 6.5 × 101 CFU/tick; but for the low inoculum dose none of the ticks were positive for LVS on day 7.
Tissue localization of LVS in A. americanum adults
To determine the dissemination of F. tularensis in tick tissues following infection by ingestion, unfed flat adult ticks were capillary fed tick meal containing 107 CFU/mL LVS and subsequently dissected on various days post-capillary feeding to determine the tissue dissemination of LVS. In adult A. americanum, LVS penetrated the gut and disseminated to the hemolymph and salivary glands within 24 hours of capillary tube feeding (Fig 6). There were no statistically significant differences in the quanta of infection between the different tissues at any time point.
Fig 6. Tissue dissemination of LVS in adult A. americanum.
Solid black bar- gut, white bar with dots- hemolymph, white bar with diagonal lines- salivary gland,. For each time point the n was 5.Error bars indicate standard deviation.
LVS secretion in infected tick saliva and infectivity in mice
To determine whether LVS disseminated to salivary glands of adult ticks is secreted into saliva and whether these secreted bacteria are infectious, partially fed A. americanum adults were first inoculated with LVS i.h., and salivary secretion induced by a secretagogue. On day 2 post i.h. inoculation, LVS was detected in gut and salivary gland and was also secreted in the saliva (Fig 7) with a mean level of 1.0±0.1 × 103CFU/μl of saliva. There were no statistically significant differences in the quanta of infection between the different tissues at any time point. In a subsequent experiment, ticks were capillary tube fed with LVS, salivation induced and saliva collected 2 days post-capillary feeding, and saliva pooled, and injected into mice to determine the ID50 for LVS in saliva. Similar to what was observed for i.h. inoculation, when ticks were capillary tube fed LVS, the bacterium disseminated from gut to hemolymph and was secreted in the saliva within two days post-capillary tube feeding (Fig 7) with a mean level of 2.0±0.8 × 102CFU/μl of saliva. The ID50 for LVS in tick saliva in BALB/c mice injected intra-peritoneally (i.p.) was <5 CFU as compared to 10 CFU for laboratory cultured LVS (Table 1). The mice that had the highest inoculation dose showed clinical signs after 3 days post-injection while the mice that had the lowest inoculation dose showed clinical signs at 5 days post-injection. All mice not showing any clinical signs were euthanized at the end of the study (7 days post injection). Bacterial culture from mouse liver and spleen showed that these tissues were heavily infected with LVS.
Fig 7. Tissue dissemination of LVS in adult A. americanum post inoculation.
Solid black bar—gut, white bar with dots–hemolymph, white bar with diagonal lines—salivary glands, and white bars with cross marks–saliva (CFU/μl of saliva). For each time point, n was 5. Error bars indicate standard deviation. For day 2 post-capillary tube feeding salivary glands were not collected.
Table 1. ID50 of F. tularensis LVS from A. americanum salivary secretions (post-capillary tube feeding) for groups A-E and laboratory cultured LVS for groups F-J in BALB/c mice.
| Group | Inoculum (CFUs of LVS) | Fraction of infected mice |
|---|---|---|
| A | 5 | 6/6 |
| B | 50 | 6/6 |
| C | 500 | 6/6 |
| D | 5000 | 6/6 |
| E | 10000 | 6/6 |
| F | 0.4 | 0/6 |
| G | 4 | 2/6 |
| H | 39 | 6/6 |
| I | 194 | 6/6 |
| J | 387 | 6/6 |
| Control | Nil ɑ | 0/6 |
ɑ Control group of mice was injected with uninfected A. americanum salivary secretions.
Transovarial transmission not observed for LVS infected gravid adult female A. americanum to larvae via the egg stage
To assess whether female A. americanum transovarially transmit F. tularensis to hatched larvae, adult females fed on sheep to repletion were inoculated one day post-repletion with LVS i.h., and dissemination of LVS to various tissues and oviposited egg masses assessed along with transovarial transmission to hatched larvae. After dissection of the infected gravid ticks at completion of oviposition, F. tularensis was detected in hemolymph, gut, Malpighian tubules, and ovaries (data not shown); but only 5/18 egg masses oviposited between days 10 and 50 post-injection were positive for F. tularensis by microbial culture (range 0.08 to 9.0 CFUs/mg egg mass). Transovarial transmission of F. tularensis to hatched larvae was not detected by either microbial culture or RT-qPCR for larvae hatched from infected and non-infected egg masses.
Discussion
The persistence of tick borne F. tularensis subspecies tularensis in the south-central United States is perpetuated by a tick-small mammalian host enzootic cycle of sylvatic tularemia [8,19,20]. Historically, D. variabilis has been considered the primary vector for F. tularensis in this endemic region [1]. However, the most abundant tick species found in this region is A. americanum, which consistently accounted for more than 90% of the total ticks in a number of tick surveys done in this region [8,9,21,22]. The host questing seasonality as well as the host feeding preference likely determines the role A. americanum may play in maintenance of the enzootic tularemia cycle as well as its potential for bridging infections to incidental hosts such as humans. In addition, the capacity of F. tularensis to maintain colonization of tick salivary glands is another factor determining the roles these tick may play as vectors for tularemia. We report herein a study of the capacity of F. tularensis to maintain colonization in A. americanum and assess the potential for A. americanum to serve as a vector of F. tularensis based on colonization capacity and natural history.
The only previous quantitative studies of F. tularensis persistence in A. americanum known to the authors were published by Hopla in 1953 and 1955. In the 1955 study, Hopla sought to characterize persistence and multiplication of F. tularensis in various stages of A. americanum infected by acquisition feeding as ticks as larvae. Our study differs from the Hopla study in that A. americanum larvae, nymphs and adults were infected with F. tularensis by artificial capillary tube feeding. An advantage of acquisition feeding studies like Hopla’s are that ticks receive physiologic cues while feeding potentially important for both the tick vector and the bacterium that are likely missing in capillary tube feeding infection of ticks such as our experiments. However, using capillary tube feeding to infect ticks, relatively large numbers of ticks can be infected by exposure to an inocula of standardized bacterial suspension, whereas with acquisition feeding, the duration and intensity of the bacteremia in F. tularensis infected laboratory animals is short and variable, respectively, making it difficult to standardize tick exposure to the agent. A recent report confirmed the difficult of acquisition feeding for infection of A. americanum with LVS [7]. In particular, these researchers observed that acquisition feeding with A. americanum on LVS infected mice was more variable and prolong than similar acquisition feeding with D. variabilis, making acquisition feeding for LVS infection of A. americanum less practical for generating sufficient numbers of infected ticks for study.
A second difference of our study and Hopla’s was the subspecies of F. tularensis used. Hopla used Bacterium tularense strain Sm [23], which was likely a fully virulent F. tularensis subspecies tularensis, the original of which we cannot identify, whereas we used an attenuated subspecies holarctica strain which is extensively characterized. It is not known whether attenuated F. tularensis strains such as LVS behave differently in tick vectors than wild-type strains, but a comparison of our study with that of Hopla’s may be instructive.
Of particular interest for our study is the potential for A. americanum to serve as an overwintering reservoir for F. tularensis in its sylvatic cycle by maintenance of colonization in nymphs which constitute the primary A. americanum stage that survives overwintering in the south-central U.S. endemic region [12]. From our study, A. americanum infected as larvae exhibited potential to maintain prolonged colonization as unfed nymphs compatible with overwintering. This observation is consistent with Hopla’s earlier observations in which he infected larvae and determined their quanta of infection through the adult stage by subsequent feeding on uninfected guinea pigs to molt infect larvae to nymphs and adults [23]. Hopla reported similar quanta of infection in engorged larvae and unfed nymphs to quanta that we observed in 1 day post-capillary feeding larvae and 14 day post-molting nymph of log 4 and 5, respectively. Hopla refed these unfed nymphs molted from infected larvae immediately following molting such that there is no comparable data for persistence of F. tularensis unfed nymphs in his study to assess potential for overwintering of the agent in unfed nymphs. From our study, we observed that F. tularensis persisted in unfed A. americanum nymphs for up to 168 days post-capillary feeding (147 days post molting), which was the longest sampling time. However, the tissue distribution of F. tularensis in unfed A. americanum nymphs changed from gut and salivary glands at 19 days after molting to primarily gut at 147 days post-molt. It would appear that F. tularensis would have to re-disseminate to salivary glands for the nymphs to be infective for initiating the next seasonal cycle. There is some evidence from Hopla’s studies and for other tick and pathogens that refeeding does elicit pathogen secretion [23–26].
While our observations for quanta of infected larvae and unfed nymphs molted from infected larvae are similar to those of Hopla, our observation for infect nymphs and adults molted from infect nymphs differ from Hopla’s infected larvae molted to nymphs and subsequently to adults [23]. In Hopla’s experiments, the infection quanta reached log 9, the highest level observed at any stage, for nymphs prior to molting to adults, but then declined to log 6.5 for unfed adults at 5 weeks post-molting. In contrast, at 21 days post-capillary feeding infection of nymphs corresponding to Hopla’s “prior to molting” stage, nymphs in our experiment exhibited a decline in infection quanta to a nadir of log 2 before rebounding at 5 weeks post-molting as unfed adults to log 4. Pre-molting nadirs of infection levels have previously been observed by us for F. tularensis in D. variabilis as well as by others for Borrelia burgdorferi in Ixodes dammini [5,27]. It is assumed that high levels of bacterial CFUs/ticks correlates with active infection and superior potential for vector transmission; however, the relationship between infection level and vector competency is not known with any specificity for tularemia.
In adult A. americanum infected as nymphs by capillary feeding, we observed that F. tularensis maintained infection in only 10% of ticks for a period up to 140 day post-molting. This is in contrast to Hopla’s observations for adult ticks infected with F. tularensis via acquisition feeding as larvae retained infection through molts to adults with 79% of these adult ticks being positive for F. tularensis at 150 day post-molting [23]. Based on these observations, A. americanum nymphs or adults acquiring infection as either larvae or nymphs could act as enzootic vectors for natural sylvatic hosts or as bridging vectors for F. tularensis of incidental hosts such as humans during spring-summer host seeking periods. In particular, the host feeding preference of nymph and adult A. americanum for larger mammalian species such as humans would be expected to facilitate it as a bridging vector for F. tularensis in humans.
As mentioned earlier for nymphs, an additional consideration for A. americanum serving as a vector of F. tularensis is secretion of the agent in saliva. For A. americanum infected as adults, F. tularensis penetrated the gut and reached the salivary glands as early as 24 hours post-capillary tube feeding in adult A. americanum. This is in contrast to our earlier report for D. variabilis, in which F. tularensis required 2 to 3 weeks to penetrate gut and reach the hemolymph and salivary glands [5]. The rapid dissemination of F. tularensis in A. americanum has implications regarding the vector competence of this tick. The extrinsic incubation period, demonstrated by the presence of F. tularensis in the saliva of the infected A. americanum adult ticks was found to be as low as 48 hours post-capillary tube feeding. This short extrinsic incubation period may have limited significance for transmission of F. tularensis by adult female A. americanum which are committed feeders, but adult male A. americanum are intermittent feeders such that adult male which acquire F. tularensis from an infected host can drop off and infest a second host and transmit F. tularensis to this new host [14], similar to the role of intermittent feeding by adult male Dermacentor andersoni ticks for the transmission of Anaplasma marginale in cattle [28,29]. However, the likelihood of adult male A. americanum acquiring F. tularensis based on their preferential feeding on larger host makes subsequent intermittent feeding transmission of F. tularensis to another large incident host such as humans appear problematic.
All three stages of A. americanum including adults, nymphs, and larvae have been found naturally infected with F. tularensis in nature [9,11,22]. The presence of F. tularensis in unfed larvae raised the possibility of transovarial route of transmission in this tick species. We examined the possible transovarial transmission of F. tularensis in A. americanum and found that occasional transmission to eggs could occur although with a relatively low level of F. tularensis infection per egg mass. However, the transmission of F. tularensis from eggs to hatched larvae was not observed. Despite the detection of F. tularensis in unfed A. americanum larvae in nature, the potential role of transovarial transmission of F. tularensis from infected adult females to hatched larvae in maintenance of F. tularensis infections in A. americanum appears unlikely. The lack of finding of large number of infected larvae in nature also supports the conclusion reached by previous researchers that transovarial transmission of F. tularensis in ticks is the exception rather than the rule [4,30].
Although extensively studied, the reservoir host for F. tularensis is still questionable in the South-Central United State endemic region [31]. The high mortality rate for tularemia in lagomorphs makes status of this species as a reservoir of F. tularensis uncertain [31]. Rodents are another obvious candidate as a reservoir host for tularemia, but there is currently insufficient data to support their role as a reservoir. Ticks are the third species in the south-central United States endemic region which may maintain F. tularensis between seasonal outbreaks. The data presented herein demonstrates the potential for A. americanum nymphs which acquire F. tularensis as larvae to maintain the agent, overwinter and to initiate new seasonal outbreaks via a tick-small mammalian life cycle. Additional studies are needed to address the issue of whether rabbits, rodents or ticks serve as reservoirs for F. tularensis in its endemic regions [20,32,33]. Based on Hopla’s and our studies, nymph or adult A. americanum which acquired F. tularensis as larvae or nymphs may serve as bridging vectors by recrudescence of F. tularensis colonization from tick gut to salivary glands during feeding to initiate subsequent seasonal tularemia epizootics.
Supporting Information
A and B- Immunostained sections of nymphs fed with LVS at 40 day post-capillary tube feeding (adsorbed serum used as primary antibody). C and D- Immunostained sections of unfed ticks using F. tularensis antiserum as primary antibody. 200x magnification.
(TIFF)
Acknowledgments
Authors thank Lab Animal Resources Facility, Oklahoma State University for providing the lab space for the mice experiments (IACUC protocol VM-10-1), Robbin Madden, Department of Entomology and Plant Pathology, Oklahoma State University for her helpful suggestions with the chemically induced salivary secretion in ticks, and the Tick Rearing Facility, Department of Entomology and Plant Pathology, Oklahoma State University for supplying the colony reared ticked used in the experiments described herein. This research was supported by the Center for Veterinary Health Sciences, Oklahoma State University.
Data Availability
All relevant data are within the paper and its Supporting Information files.
Funding Statement
The study was funded by the Center for Veterinary Health Sciences, Oklahoma State University.
References
- 1. Eisen L. A call for renewed research on tick-borne Francisella tularensis in the Arkansas-Missouri primary national focus of tularemia in humans. J Med Entomol. 2007; 44: 389–397. [DOI] [PubMed] [Google Scholar]
- 2. Goodman JL, Dennis DT, Sonenshine DE. Tick-Borne Diseases of Humans. Washington, D.C.: ASM press; 2005. [Google Scholar]
- 3. Jellison WL. TULAREMIA in North America. Missoula, Montana: University of Montana; 1974. [Google Scholar]
- 4. Bell JF. Infection of ticks (Dermacentor variabilis) with Pasteurella tularensis . J Infect Dis. 1945;76: 83–95. [Google Scholar]
- 5. Mani RJ, Reichard MV, Morton RJ, Kocan KM, Clinkenbeard KD. Biology of Francisella tularensis Subspecies holarctica Live Vaccine Strain in the Tick Vector Dermacentor variabilis . PLoS One. 2012;7: e35441 10.1371/journal.pone.0035441 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 6. Philip CB, Jellison WL. The American Dog Tick, Dermacentor variabilis as a Host of Bacterium tularense . Public Health Rep. 1934; 49: 386–392. [Google Scholar]
- 7. Coburn J, Maier T, Casey M, Padmore L, Sato H, Frank DW. Reproducible and Quantitative Model of Infection of Dermacentor variabilis with the Live Vaccine Strain of Francisella tularensis . Appl Environ Microbiol. 2015;81: 386–395. 10.1128/AEM.02917-14 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 8. Hopla CE. The transmission of tularemia organisms by ticks in the southern states. South Med J. 1960; 53: 92–97. [DOI] [PubMed] [Google Scholar]
- 9. Calhoun EL. Natural occurrence of tularemia in the lone star tick, Amblyomma americanus (Linn.), and in dogs in Arkansas. Am J Trop Med Hyg. 1954; 3: 360–366. [DOI] [PubMed] [Google Scholar]
- 10. Hopla CE. Experimental studies on tick transmission of tularemia organisms. Am J Hyg. 1953; 58: 101–118. [DOI] [PubMed] [Google Scholar]
- 11. Hopla CE, Downs CM. The isolation of Bacterium tularese from the tick Amblyomma americanum . J Kansas Entomol. 1953; Soc 26: 72–73. [Google Scholar]
- 12. Kollars TM Jr, Oliver JH Jr, Durden LA, Kollars PG. Host association and seasonal activity of Amblyomma americanum (Acari: Ixodidae) in Missouri. J Parasitol. 2000; 86: 1156–1159. [DOI] [PubMed] [Google Scholar]
- 13. Kollars TM Jr, Oliver JH Jr, Masters EJ, Kollars PG, Durden LA. Host utilization and seasonal occurrence of Dermacentor species (Acari:Ixodidae) in Missouri, USA. Exp Appl Acarol. 2000; 24: 631–643. [DOI] [PubMed] [Google Scholar]
- 14. Sonenshine DE. Biology of Ticks. New York: Oxford University Press; 1991. [Google Scholar]
- 15. Maier TM, Havig A, Casey M, Nano FE, Frank DW, Zahrt TC. Construction and characterization of a highly efficient Francisella shuttle plasmid. Appl Environ Microbiol. 2004; 70: 7511–7519. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 16. Versage JL, Severin DD, Chu MC, Petersen JM. Development of a multitarget real-time TaqMan PCR assay for enhanced detection of Francisella tularensis in complex specimens. J Clin Microbiol. 2003; 41: 5492–5499. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 17. Jaworski DC, Simmen FA, Lamoreaux W, Coons LB, Muller MT, Needham GR. A secreted calreticulin protein in ixodid tick (Amblyomma americanum) saliva J Insect Physiol. 1995; 41: 369–375. [Google Scholar]
- 18. Lennette EH. General principles underlying laboratory diagnosis of viral and rickettsial infections; Lennette EH, Schmidt NJ, editors. New York: American Public Health Association Inc; 1964. [Google Scholar]
- 19. Assal NR, Lindeman RD, Carpenter RL. Epidemiologic study on reported human tularemia in Oklahoma, 1944–65. J Okla State Med Assoc. 1968; 61: 120–124. [PubMed] [Google Scholar]
- 20. Petersen JM, Mead PS, Schriefer ME. Francisella tularensis: an arthropod-borne pathogen. Vet Res. 2009; 40: 7 10.1051/vetres:2008045 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 21. Brown HE, Yates KF, Dietrich G, Macmillan K, Graham CB, Reese SM, et al. An acarologic survey and Amblyomma americanum distribution map with implications for tularemia risk in Missouri. Am J Trop Med Hyg. 2011; 84: 411–419. 10.4269/ajtmh.2011.10-0593 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 22. Calhoun EL, Alford HI Jr. Incidence of tularemia and Rocky Mountain spotted fever among common ticks of Arkansas. Am J Trop Med Hyg. 1955; 4: 310–317. [DOI] [PubMed] [Google Scholar]
- 23. Hopla CE. The multiplication of tularemia organisms in the lone star tick. Am J Hyg. 1955; 61: 371–380. [DOI] [PubMed] [Google Scholar]
- 24. Piesman J, Mather TN, Sinsky RJ, Spielman A. Duration of tick attachment and Borrelia burgdorferi transmission. J Clin Microbiol. 1987; 25: 557–558. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 25. Spencer RR, Parker RR. Rocky Mountain spotted fever: infectivity of fasting and recently fed ticks. Public Health Reports. 1923; 38: 333–339. 19314866 [Google Scholar]
- 26. Katavolos P, Armstrong PM, Dawson JE, Telford SR III. Duration of tick attachment required for transmission of granulocytic ehrlichiosis. J Infect Dis. 1998; 177: 1422–1425. [DOI] [PubMed] [Google Scholar]
- 27. Piesman J, Oliver JR, Sinsky RJ. Growth kinetics of the Lyme disease spirochete (Borrelia burgdorferi) in vector ticks (Ixodes dammini). Am J Trop Med Hyg. 1990; 42: 352–357. [DOI] [PubMed] [Google Scholar]
- 28. Kocan KM, Stiller D, Goff WL, Claypool PL, Edwards W, Ewing SA, et al. Development of Anaplasma marginale in male Dermacentor andersoni transferred from parasitemic to susceptible cattle. Am J Vet. 1992; Res 53: 499–507. [PubMed] [Google Scholar]
- 29. Kocan KM, Goff WL, Stiller D, Edwards W, Ewing SA, Claypool PL, et al. Development of Anaplasma marginale in salivary glands of male Dermacentor andersoni . Am J Vet Res. 1993; 54: 107–112. [PubMed] [Google Scholar]
- 30. Hopla CE. The ecology of tularemia. Adv Vet Sci Comp Med. 1974; 18: 25–53. [PubMed] [Google Scholar]
- 31. McCahan GR, Moody MD, Hayes FA. An epizootic of tularemia among rabbits in northwestern South Carolina. American Journal of Epidemiology. 1962; 75: 335–338. [Google Scholar]
- 32. Petersen JM, Molins CR. Subpopulations of Francisella tularensis ssp. tularensis and holarctica: identification and associated epidemiology. Future Microbiol. 2010; 5: 649–661. 10.2217/fmb.10.17 [DOI] [PubMed] [Google Scholar]
- 33. Kugeler KJ, Mead PS, Janusz AM, Staples JE, Kubota KA, Chalcraft LG, et al. Molecular epidemiology of Francisella tularensis in the United States. Clin Infect Dis. 2009; 48: 863–870. 10.1086/597261 [DOI] [PubMed] [Google Scholar]
Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.
Supplementary Materials
A and B- Immunostained sections of nymphs fed with LVS at 40 day post-capillary tube feeding (adsorbed serum used as primary antibody). C and D- Immunostained sections of unfed ticks using F. tularensis antiserum as primary antibody. 200x magnification.
(TIFF)
Data Availability Statement
All relevant data are within the paper and its Supporting Information files.