Lyme Disease in Clinically Sick Dogs in Southern US: Assessing Geographical and Seasonal distribution during year 2012 – 2011 in Texas

Research Article

Lyme Disease in Clinically Sick Dogs in Southern US: Assessing Geographical and Seasonal distribution during year 2012 – 2011 in Texas

*Corresponding author: Dr. Maria D. Esteve-Gassent, Department of Veterinary Pathobiology, VMA316, College of Veterinary Medicine and Biomedical Sciences , TAMU-4467 College Station, TX-77843-4467 Phone: 979-845-1117. Fax: 979-862-2344  Email:


Lyme disease is not reported in veterinary medicine, even though dogs have been described as sentinels for this disease. Most of the canine Lyme disease studies in the US have been focused in endemic areas, and few studies have been done in Southern US. The objective of this study was to evaluate the seroprevalence of Lyme disease in dogs suspected of having a tick borne illness in Texas. A total of 831 dog-serum samples collected during 2011-12 were tested. Ticks collected though passive surveillance were also evaluated for the presence of Borrelia burgdorferi and B. lonestari. Results showed that B. burgdorferi sero-reactive canine samples occurred from November through February, which also coincided with peak detection of B. burgdorferi in ticks. In contrast, zero B. lonestari infected ticks were detected. These results suggest that the levels of B. burgdorferi sero-reactivity in sick dogs found in Texas is similar to that observed in other non-endemic states. Canine Lyme disease season in Texas was observed to extend from fall to early spring, which coincides with the active season for the adult tick vector Ixodes scapularis. In addition, ticks infected with B. lonestari might be less prevalent in Texas than previously speculated.

Keywords: Borrelia burgdorferi; Borrelia lonestari; ELISA; Canine Lyme Disease; Tick Borne Diseases; P66; dogs; geographic distribution


Lyme disease (LD) is the most prevalent arthropod-borne infection in the United States [1, 2]. A significant increase in the number of human reported cases has been observed in the past few years, classifying LD as a re-emerging infection. Borrelia burgdorferi, the causative agent of LD, is transmitted to humans and companion animals through the bite of infected Ixodes ticks [3, 4]. Lyme borreliosis is therefore an important public health issue, particularly in endemic areas. Canine LD occurs in the majority of infected animals in a subclinical status [5, 6], and in some endemic areas the majority of LD positive dogs do not develop any signs of disease [5, 6]. When becoming ill, the syndrome commonly associated with canine LD includes polyarthritis, lymphadenopathy and glomerulonephritis [7, 8].

Since canine LD is not a reportable disease, the epidemiological data available to estimate the number of confirmed animal cases nationwide is incomplete [9]. Scant epidemiological data is available from nationwide studies carried out by Bowman et al. [9], Qurollo et al. [10], the IDXX dogs and ticks interactive maps (http://www.dogsandticks. com/diseases_in_your_area.php ) and the Banfield Annual report on The State of Pet Health™ 2014 report [11]. Detection of antibodies to the spirochete B. burgdorferi in these studies was achieved using commercially available kits [9, 10].  Even considering these sources, very little has been done in southern US. In Texas in particular, the IDXX interactive map describes 2,684 veterinary LD cases; 6,714 Ehrlichiosis; 65,058 Heartworm and 3,344 Anaplasmosis cases reported in the last 5 years (last checked May 4th 2016). In 2010, out of 500 canine LD tests performed at the Texas A&M Veterinary Medical Diagnostic Laboratory (TVMDL), 40% of the positive results came from Texas, while the other 60% came from out of state (Sandy Rodgers, personal communication). The majority of the LD positive animal cases came from East Texas. When analyzing the confirmed number of human cases in Texas over the last 10 years, the region with highest incidence of human LD is the Cross Timbers eco-region located in central Texas [12]. There have also been several human cases reported in West Texas and near the Panhandle, where very few animal cases have been detected. The lack of information and awareness of LD in rural areas across Texas coupled with a better understanding of LD in the metropolitan areas could explain the detection discrepancies between the two. Consequently, surveillance programs could estimate the seroprevalence for B. burgdorferi infection in wildlife and the proportion of ticks carrying B. burgdorferi DNA, and thereby determining which areas pose a risk for LD in Texas [13-18].

Currently, there are no reliable tests to detect the causative agent of LD in dogs or humans [19]. Nevertheless, there are various laboratory-testing methods employed in the diagnosis of LD, including serological tests such as ELISA [8, 20-22] (enzyme immunoassay), IFA (immunofluorescentantibody assay), Western Blot (immunoblot, IB) [19] and multiplex ELISA [23]. In both human and veterinary medicine, the commercially available serological diagnostic kits are expensive and mostly detect chronic cases with clinical signs, but are not efficient in detecting acute and/ or subclinical cases [23-28]. PCR has been proven to be the most effective of these methods [19, 27, 29], however, it is not used as often as the serological methods because biopsy tissue rather than blood must be tested in order to gain reliable information. On the other hand, ELISA and IB have very similar testing sensitivities, and therefore these two methods are the most commonly used in diagnostic laboratories. The LD serological diagnostic techniques mostly used in veterinary medicine are the IFA, using whole cells, and the C6 ELISA tests, which utilizes the C6 epitope of the variable surface antigen VlsE [30-40]. In addition, the IFA is known for subjective interpretation and low sensitivity, while the ELISA test is known to have high sensitivity [34, 41, 42]. Therefore, several companies and research groups are currently developing new and improved methods that are more specific and sensitive than the methods mentioned above [9, 23, 43, 44].

Companion animals such as dogs and horses have been considered in previous studies as good sentinels for LD in endemic areas of the US [9, 16, 45, 46].  Due to the sparse information available in the literature regarding canine LD in Southern US, our team took the challenge to evaluate the B. burgdorferi sero-reactivity present in ailing dogs from Texas suspected of a tick-borne illness. This study was performed in collaboration with diagnosticians from TVMDL. To this end, dog serum samples were collected from October 2011 through October 2012 and evaluated [47, 48]. Thus, the objectives of this study are: i) to determine the temporal distribution of LD sero-reactivity in dogs with clinical illness in the State of Texas; ii) assess the cross-reactivity of the tests of choice with other spirochetal bacteria, in particular Leptospira interrogans, commonly found in Texas, and iii) evaluate the location of canine serum samples sero-reactive to B. burgdorferi in the State of Texas and how it compares with both the distribution of the tick vector I. scapularis, and the presence of human LD cases in the same time frame.

Materials and Methods


A total of 924 canine serum samples were transferred from the Texas A&M Veterinary Medical Diagnostic Laboratory (TVMDL) to the College of Veterinary Medicine & Biomedical Sciences at Texas A&M University after the 15-day legal hold period, in accordance with the Material Transfer Agreement between both institutions.  No confidential information regarding the pet owners and/or veterinary clinic where the animals were evaluated was provided. The experiments described in this study were conducted under the Institutional Biosafety Permit number 2010-036 and 2013-039.

Canine Serum Samples

All serum samples were obtained from individual animals suspected of having a tick borne illness during October 2011 to October 2012. The information obtained from TVMDL for each sample included the case number, county of origin, zip code, the sample arrival date, the IFA analysis completion date, and the resultant antibody titer and IFA test results. All samples were tested at TVMDL for LD utilizing standardized IFA tests (Focus Diagnostics, Inc.). Thirteen additional negative control samples were obtained from a colony of dogs housed at Texas A&M University, thus the total sample size was 937 specimens. Samples obtained at different times of the year came from different animals (i.e., not repeated measurements were taken). One hundred and six samples were found in bad conditions and were eliminated from the study, leaving a total of 831 samples to be studied.

Borrelia burgdorferi Strains and Growing Conditions

b.burgdorferi B31 A3 virulent isolate was used for the generation of antigens for the ELISA test used throughout this study. In order to obtain an antigen profile similar to the one observed during natural infection, this bacterium was grown at room temperature and pH 7.6 to mimic the unfed tick conditions. Once the bacterial cultures reached a cell density of 1-2×107 spirochetes/ml a subculture was transferred to 37ºC and pH 6.8 mimicking the conditions in the tick upon feeding [49]. After cultures reached a cell density of 3-5×107 spirochetes/ml cells were harvested, washed three times with HBSS buffer (HyClone, Thermo Scientific, Inc. Pittsburgh, PA), quantified, and lysed using 0.1mm glass beads in 2 ml screw cap tubes in a BeadRuptor 24 (Omni Internationa, Inc. Kennesaw, GA). After the lysis cycle, glass beads were sedimented by quick centrifugation, and the supernatants were stored at -20ºC, in 1ml aliquots containing 109 spirochete/ml, until use in the ELISA assays. rP66 Protein Expression and Purification

The codding region of P66 (BB0603) was PCR amplified utilizing the forward primer 5’-ACGCGCTAGCATGAAAAGCCATATTTTA-3’ and the reverse primer 5’- ACGCCTCGAGGCTTCCGCTGTAGGCTAT-3’ and then cloned as a fusion protein with His*tag in the expression vector pET23a (Novagen ® EMD Millipore Coorporation, Germany). NheI and XhoI restriction sites (underlined and bold primer sequence above) were used for cloning in pET23a, (pLE32B) prior to transformation into E. coli RosettaTM strain for expression (Novagen ® EMD Millipore Coorporation, Germany) (LE32, Table 1) [49-51]. To purify rP66, an E. coli RosettaTM culture containing the pLE32 construct was started in LB broth medium containing ampicillin 100 µg/ml (Amp100) and chloramphenicol 20 µg/ml (Can20) at 37ºC and agitated overnight. A 1:100 dilution of the overnight culture was used to start a oneliter culture in LB broth containing Amp100 and Cam20. Large cultures were agitated for 4-5 hours at 37ºC or until reaching an OD600nm of 0.5 to 0.8. The expression of the recombinant protein was induced by adding Isopropyl β-D-

1-thiogalactopyranoside (IPTG) at a final concentration of 1 mM to the cultures followed by continual agitation for 2 hours at 37°C. After the 2-hour induction process, cells were harvested by centrifugation for 10 minutes at 10,000 rpm and 4ºC and pellets were stored at -80ºC until use [52].

Table 1. Bacterial strains and plasmids utilized in this study

After thawing on ice, the pellets of RosettaTM cells expressing rP66 were re-suspended in 25 ml of Lysis Buffer (50mM sodium phosphate, 8 M urea, 300 mM NaCl, 20 mM imidazole pH 7.4) containing 100µl of the protease inhibitor HaltTM cocktail (Thermo Scientific, Inc. Pittsburgh, PA). Cells were disrupted by French Press (Thermo Scientific, Inc. Pittsburgh, PA) and lysates were centrifuged for 10 minutes at 10,000 rpm and 4ºC to remove debris as previously described [49]. Supernatants were mixed with 5 ml of Nickel beads (His60 Superflow™ resin Clonetech Laboratories, Inc. Mountain View, CA) equilibrated with Wash Buffer (50mM sodium phosphate, 8 M urea, 300 mM NaCl, 40 mM imidazole pH 7.4) and incubated overnight at 4°C with continuous gentle agitation. After overnight binding, beads were cleaned by centrifugation for 5 minutes at 500 rpm and 4ºC to remove any unspecific binding. Beads with the recombinant His-tag containing proteins were allowed to pack at room temperature in a chromatography column (BioRad Laboratories, Inc. Hercules, CA). Recombinant P66 was eluted by adding 10 ml of the elution buffer (50mM sodium phosphate, 8 M urea, 300mM NaCl, 300mM imidazole pH 7.4) at a time to the column. A total of approximately 20 ml of elution buffer (2 runs of 10 ml) was needed to ensure the adequate purification of the rP66. Five hundred microliters elution fractions were collected and saved at -80°C until use. In order to determine which elution fractions contained the most recombinant proteins, an aliquot of the induction pellet, post-absorption supernatant, wash fractions and elution fractions were separated in a 12% SDS-PAGE gel. Proteins on the gels were visualized by Coomassie Brilliant Blue staining.

The fractions containing the recombinant proteins were cleaned and concentrated using the Amicon® filtration system (EMD Millipore Corporation, Germany) with a cut off pore size of 10kDa to ensure the retention of the protein of interest [49]. The concentrated protein was further cleaned by dialysis to ensure the elimination of the denaturing agent urea and Imidazol used to elute the protein during the purification steps. To this end, the concentrated protein was injected into a dialysis cassette (Slide-ALyser® Cassette, Thermo Scientific, Inc. Pittsburgh, PA) and incubated in 500 ml of dialysis buffer (50mM sodium phosphate, 300mM NaCl) for 2 hours at room temperature while being continuously stirred (following manufacturer’s recommendations). After 2 hours, the concentrated fractions were recovered and stored at -80°C. Aliquots were used to determine protein concentration by the BCA assay (Thermo Scientific, Pittsburgh, PA).

ELISA (Enzyme linked immunosorbed assay)

Two different ELISA tests were performed to determine the level of reactivity of dog sera samples to B. burgdorferi, whole cell lysates (aBb), and the rP66 [49]. Out of the 937 total serum samples collected (including the negative controls), 831 canine serum samples that were in good conditions were analyzed by both Bb and rP66 ELISA tests. To this end, 96-well plates (Nunc MaxiSorb®, Thermo Scientific, Inc. Pittsburgh, PA) were coated with the purified rP66 at a concentration of 500ng/well or with 107 cells/ well for the B. burgdorferi lysate in carbonate buffer pH 9.4 and then incubated at 4ºC overnight as previously described [52]. After coating, plates were washed three times with Phosphate Buffer Saline containing 0.5% Tween 20 (PBS-T). Plates were then blocked with 200μL of PBS-T containing 3% BSA at 4ºC overnight. After blocking, plates were washed three times in PBS-T and incubated for 1 hour with 1:400 dilution of each of the animal samples in PBS-T containing 1% BSA. Blank wells were incubated with buffer only. After washing the plates 3 times with PBS-T, a 1:2,000 dilution of the anti-dog HRP-conjugated IgG antibody (Rockland Immunochemicals, Gilbertsville, PA) was added to each well. The plates were incubated for 1 hour at room temperature followed by 3 washes in PBS-T and then incubated with 100µl/well of o-phenylenediamine dihydrochloride color substrate (Thermo Scientific, Inc. Pittsburgh, PA) for 30 minutes in the dark. Plates were read at a wavelength of 450nm using a plate reader and software (BMG LABTECH OMEGA, Germany).

Western Immunoblot (IB)

Two hundred and thirty two dog sera samples were evaluated by IB using a commercially available human B. burgdorferi strip system (Trinity Biotech, B. burgdorferi Marblot™ Strip Test System, Ireland), adapted for testing dog samples. Samples were selected based on their reactivity in ELISA and IFA tests. Briefly, one pre-coated strip with B. burgdorferi antigens was used per dog serum sample. For each sample or control strip (Positive and Negative), a channel in a 12-strip plate was filled with 2 ml of 1X sample diluent/wash solution provided by the manufacturer. After strips were equilibrated for 5 minutes, 20 μL of each of the samples was added to the appropriately marked channel and incubated at room temperature for 30 minutes. Strips were washed three times by adding 2 mL of sample diluent/wash solution to each channel of the strip incubation tray and incubated for 5 minutes with vigorous agitation. Two ml of a 1:2,000 dilution of the anti-dog alkaline phosphatase conjugated IgG antibody (Rockland Immunochemicals, Gilbertsville, PA) was added to each strip containing well, and incubated for 30 minutes at room temperature. Strips were then washed 3 times and 2 mL of color developing solution was added to each channel. All strips were incubated for 6 minutes to allow color development. Strips were then washed with 2 mL of deionized water, air-dried, and evaluated. The presence or absence of the following 13 bands was then recorded: a93, a66, a60, a58, a45, a41 (Flagella), a39 (BmpA), a34, a31 (OpsA), a30, a28, a23 (OpsC) and a18.

Leptospira cross-reactivity

A random subset of 91 canine serum samples were tested for various Leptospira serovars prevalent in Texas, namely Autumnalis, Bratislava, Canicola, Grippotyphosa, Ichterohaemorrhagiae, Pomona and Sejroe through the TVMDL laboratory. The Microscopic Agglutination Test (MAT) was used to evaluate each one of the samples submitted as per the ACVIM small animal consensus statement [53]. Antibody titers to each of the positive serovars were determined for each of the positive samples tested.

Tick collection and Borrelia burgdorferi detection

To detect the prevalence of Borrelia burgdorferi infection in the tick population, ticks were submitted to our laboratory from three Texas Parks and Wildlife management areas, animal shelters and veterinary clinics [54]. In total, we received ticks from 20 counties in the State of Texas. Upon arrival, ticks were identified using dichotomous keys [5558] and processed for DNA extraction and B. burgdorferi detection as previously described [54].  Location and infection rate of the Ixodes scapularis ticks used in this study were presented in the study by Feria-Arroyo et al. [54]. The sequences of positive ticks were published elsewhere [54, 59]. In addition, in order to detect the potential presence of B. lonstari in Amblyomma ticks, all Amblyomma spp. ticks collected during the same time frame were tested for the presence of relapsing fever Borrelias utilizing glpQ, and broadly reactive flaB specific primers as previously described [60]. No ticks were collected from the dogs analyzed by serology in this study. Therefore, all tick samples and serum samples were collected independently, to avoid potential biases.

Geographic Information System (GIS)

Spatial analysis of the data was conducted using ArcGIS 10. The mapping of seropositive canine samples was performed considering the different eco-regions in Texas. After acquiring the ZIP code of each sample, each was given a centroid and the centroids were mapped in regards to the eco-regions in which they were located.

Statistical Methods

Continuous variables were described by median and interquartile range (IQR), and groups compared using the non-parametric Kruskal-Wallis test. The number and percent positive described the dichotomous variables and groups compared using Fisher’s exact test.

All subsets of the six IB bands 18, 23, 30, 34, 41 and 93, were considered as possible diagnosis sets, and for each IB band the values (0 = absent, 0.5 = weak, 1 = present) were summed, and then different cut off-points for diagnosis were considered (higher than cut off-point = positive, lower than cut off-point = negative). At the same time, various cut offpoints based on combinations of the two ELISAs OD450nm tests were also considered. Based on higher sensitivity and specificity of IB in human diagnostics, the preferred cut off value was that which gave highest average of sensitivity and specificity of ELISA diagnosis compared to the IB diagnosis (Additional information 2). Statistical significance was determined at the 5% level. All data handling and statistical analysis was performed in Stata MP version 12 (STATA Corp, College Station, TX) [61].


A total of 937 dog sera were evaluated and IFA tested by TVMDL. Of these, 106 samples were in bad conditions and removed from the study. From the remaining 831 samples, 219 samples and 13 control samples were analyzed by commercially available IB assay and all 831 samples were analyzed using the in-house ELISA assay. All controls were negative for IFA, and showed very low sero-reactivity in the ELISA test (aBb OD450nm < 0.350 and rP66 OD450nm < 0.650) at 1:400 dilution. Of the subset of samples tested by IB and ELISA, 125 samples tested negative for the IFA, leaving 94 IFA positive samples. Descriptive statistics are reported by group in Table S1. We divided the samples into control, meaning no possibility of exposure to LD pathogen, and corresponding to samples collected from the colony dogs, and non-control meaning dogs sera acquired through TVMDL and that were potentially exposed to the pathogen.

Presence of IB Antigen bands and ELISA Optical Densities (OD450nm) compared to IFA diagnosis

A total of 232 canine serum samples were subjected to ELISA and IB testing: control (n=13), non-control and IFA negative (n=125), and non-control and IFA positive (n=94). The proportion of samples showing each of the 13 IB antigen bands varied between the three groups. Interestingly, the 23 (ospC), 45, 58 and 66 kDa bands occurred in the control group, of which the 58 and 45 kDa occurred more often in the control than the non-control groups (see supplemental Table S1 and Additional information 1, Figure S1).

Table 2 shows average OD450nm for both ELISA tests (aBb and rP66), from 232 canine serum samples, by IB antigen band presence. P-values were obtained from t-tests of ELISA OD450nm by IB antigen band presence. For both ELISAs, p-values for the 58, 60 and 66 kDa bands were large, indicating a lack of correlation between ELISA ODs and the presence of these IB bands. The best IB antigen bands the purpose of discrimination between high and low ELISA OD450nm values were the 18, 23, 30, 34, 41 and 93 kDa bands for These data are shown in Figure 1 with 95% confidence intervals.

Determination of IB positive and ELISA positive diagnosis for LD

In order to determine which samples were positive for LD, a two-tier system similar to human LD diagnosis was utilized. In this case, IB test was used to confirm exposure to the Lyme disease pathogen. As described above, out of the 8 bands detectable in the IB assay, a subset of six IB bands 18, 23, 30, 34, 41 and 93 kDa, was considered as possible diagnosis set. For each band the values 0 = absent, 0.5 = weak, 1 = present, were summed and different cut off-points for diagnosis were considered (higher than cut off-point = positive, lower than cut off-point = negative).  Also, the use of a 0.5 value for weak bands in the IB did not provide any improvement in the concordance of the IB and ELISA tests. Therefore, weak bands can be considered negative without altering the results. Four of these six bands, 18, 23, 30 and 93kDa, were found to consistently out-perform other subsets of bands. Therefore these bands were considered in further analysis.

Using the IB as the gold standard, various cut off-lines for the two ELISA tests evaluated were considered. Within samples with low anti-B. burgdorferi (aBb) OD values (aBb OD450nm < 0.6), 76% (69/91) of samples had fewer than 2 of the IB bands present (18, 23, 30 and/or 93). On the other hand, samples with high aBb OD values (aBb OD450nm > 1.3), 75% (42/56) had 2 or more of these IB bands present. For intermediate aBb OD450nm values, utilizing the additional information from the rP66 ELISA test was helpful. Visual inspection of the scatter plot of the ELISA values for aBb verses rP66 revealed that 71% (20/28) of samples with all 4 IB bands present had rP66 ELISA values greater than those obtained using the aBb ELISA. Thus we defined ELISA positive based on the maximum of two sloped lines (Figure S2A, Additional information), representing the interaction of both ELISA tests.

Cross-reactivity – Comparison of IB and ELISA diagnosis to Leptospirosis diagnosis

Ninety-one samples were tested for leptospirosis at TVMDL and were all tested by both IB and ELISA for Lyme disease. Of the 4 IB positives, 3/4 (75%) were positive for leptospirosis, and of the 87 IB negatives, 68/87 (78%) were positive for leptospirosis. There was no significant difference between these percentages according to a Fischer’s exact test (P-value=1).

Nine of the 14 ELISA positives (64%), were positive for leptospirosis, and 62 of the 77 ELISA negatives, (81%), were positive for leptospirosis. There was also no significant difference between these percentages according to a Fischer’s exact test (P-value=0.181). Ten of the 14 ELISA positives were IB negative, and of these 6/10 (60%) were positive for leptospirosis, and of the 4 ELISA and IB positives, 3/4 (75%) were positive for leptospirosis. There was no significant difference between these percentages according

* indicates t-test significant at the   %5 level

** indicates t-test significant at the %1 level

Table 2. Mean ELISA OD450nm values (and count (n)) for both aBb and rP66 tests, from 232 canine serum samples, by results of IB antigen bands, with P-values from t-tests of ELISA OD450nm by IB antigen band presence.


to a Fischer’s exact test (p-value=1).

Thus, these samples do not provide any evidence of crossreactivity between leptospirosis MAT test and the Lyme IB and ELISA tests. In addition, the 3 samples positive for Leptospira and LD by ELISA and IB assay had low Leptospira titers to serovars Autumnalis, Bratislava, Canicola, Gippotyphosa, and Pomona, ranging from 100 to 800. The samples with high antibody titers to Leptospira serovars (from 1,200 to 12,800) were negative to B. burgdorferi by IB test. Notice that the initial dilution utilized in the ELISA test is 1:400 and 1:100 for the IB test.

Temporal and geographic distribution of the canine LD in Texas

Of the 924 non-control samples analyzed, 847 originated in

Texas. Of these, 741 were ELISA tested and 202 were also IB tested. The 106 Texas samples not tested were found in bad condition and eliminated from the study (evaporated, coagulated, etc.). A box plot of the aBb and rP66 ELISA ODs and the number of IB bands, 18, 30 and 93kDa, by month tested, is shown in Figure 2. The notable feature of this graph is the uptick in ELISA ODs and the presence of IB bands during November, December and January. The median sum of IB band strength was at least two in these months. November and December saw the highest average aBb and rP66 ODs. The averages, counts and percentages for these

Figure 1. Comparison of ELISA and Immunoblot assay (IB) for 13 control, 125 non-control and IFA negative, 94 non-control and IFA positive samples, and all groups combined. (A) Average ELISA aBb ODs, with %95 confidence intervals, by IB Antigen band presence/absence, (B) Average rP66 ODs, with %95 confidence intervals, by IB Antigen band presence/absence. IB assay bands are represented in the y-axis

Figure 2. Box plot of aBb and rP66 ELISA ODs, and the sum of the presence of IB bands 18, 30 and 93kDa, for non-control Texan canines, by month tested: October 2011-September 2012. Sample sizes: 707 (110 counties) for ELISA and 183 for IB (65 counties).data are listed in Table 2, together with results from the IFA tests, by month.

Figure 3 reveals seasonal variation in B. burgdorferi seroreactivity in Texas dog samples according to IB, ELISA and IFA using positive cutoffs described earlier. In particular, dogs were mostly sero-reactive during fall and winter months according to both the IB test and the ELISA test.

Considering IB testing as the gold standard, IFA false positives, compared to ELISA (n=741), were most common in March, June, July and August, and we noted that June, July and August were months that received the largest number of samples for LD testing at TVMDL in 2012.

Figure 3. Percent of positive cases by month according to IB, ELISA and IFA tests for LD for non-control Texan canines, by month of year tested: October 2011-September 2012. IB positive was the presence of IB bands 18, 30 and 93kDa. ELISA positive was aBb>2.35-1.65rP66 and aBb>0.07+0.8rP66. Sample size: n=183 in 65 Texas counties.

Table 3. Descriptive statistics of ELISA, IB, IFA tests for LD for serum samples from 828 Texan canines from 120 counties, by month of year tested: October 2011-September 2012. IB positive was the presence of IB bands 18, 30 and 93kDa. ELISA positive was aBb>2.35-1.65rP66  and aBb>0.07+0.8rP66.

Figure 4. Geographic localization of canine LD in Texas during the year 2011-2012. (A) Counties from which canine serum samples were received are marked in grey, while those from which seropositive samples were detected are highlighted in black. (B) Eco-regions in which the majority of canine seropositive cases were detected. Numbers indicate the percent of positive cases detected in each eco-region considering the total number of positive cases. In parenthesis is represented the percent of positive samples considering the number of submitted samples in each eco-region. Only eco-regions with more that 10% of the total positive samples were included in this map.

In addition to the temporal distribution, the geographical distribution of the seropositive dogs was evaluated by means of geographic information system (GIS). As shown in Figure 4 (supplemental table S2), a total of 110 counties out of 254 counties of the State of Texas were represented in this study. Most of the animals were from counties located in Eastern Texas (Figure 4A). From those, 38 counties showed seropositive canids (Figure 4A). When overlaying the counties from which seropositive canine samples were submitted and the eco-regions in Texas, we observed that 85.90% of the positive samples were distributed in the 5 most Eastern eco-regions (Cross Timbers, East Central Texas Plains, South Central Plain, Texas Blackland Prairies, and the Western Gulf Coast Plain, Figure 4B). Very low seropositive samples were detected in other eco-regions in Texas such Arizona/New Mexico Mountains, Central Great Plains, Chihuahua dessert, High Plains, South Texas Plains/Interior plains and Hills with xerophytic shrub and oak forest, and the Southwestern Tablelands.

Temporal distribution of ticks in Texas

OOf the 544 ticks collected in this passive surveillance effort, 13.23% were questing adults I. scapularis ticks (Table 4). No I. scapularis nymphs were submitted. Adults and nymphal stages of Amblyomma americanum, A. cajennense, Dermacentor albipictus, D. variabilis and Rhipicephalus sanguineus were also submitted. When evaluating the distribution of the different species of ticks throughout the year, the majority of the specimens were received during the months of December to March (Figure 5A). Notice that adult I. scapularis was mainly active during the months of November through February, and no specimens were submitted during the summer months. These results coincide with the period when most of the infections were detected in dogs by means of ELISA and Immunoblot assay. On the other hand, Amblyomma species and R. sanguineus ticks were received year around. Finally, D. albipictus, also known as the winter tick in Southern US, was mostly active during the months of December through February. When the I. scapularis tick specimens were tested for the presence of B. burgdorferi (Figure 5B), 45% of the ticks were positive by PCR (sequence data published elsewhere [54, 59]). In addition, none of the 75 A. americanum and 39 A. maculatum specimens were positive for glpQ or flaB specific relapsing fever Borrelial PCR markers. Therefore, we could not verify the presence of the relapsing fever Borrelia, B. lonestari, in the 114 Amblyomma spp. ticks tested from Texas.

Figure 5. Temporal distribution of total number of ticks per species received each month during the year of study (A). (B) Temporal distribution of I. scapularis tick vector positive for B. burgdorferi.

* Percent of the total ticks collected per species is represented in  parenthesis.

Table 4. Ticks collected by passive surveillance in the State of Texas during October 2011 through October 2012.


Commercially available diagnostic tests for canine LD include IFA and multiplex ELISA, which vary in pricing and sensitivity. Additionally, LD is not a reportable disease in veterinary medicine, which makes the selection and standardization of a test a challenging task. Here, we evaluated the immune reaction of Texan dogs suspected of a vector borne infection using different in-house and commercially available Lyme testing methods: IFA, ELIA and IB respectively. The tests were chosen based on what is currently offered as a Lyme test in common veterinary practices in the US and Europe [8, 9, 35, 45, 46, 62, 63]. During the present study, the results obtained with the in-house ELISA and the commercially available IB assay were in agreement more often with percentage of positive dogs of 9.85% and 10.40% respectively, while IFA tended to overestimate the number of positive cases at 14.76%. This result highlights the potential subjectivity of the IFA assay and its interpretation, mostly due to the interjection of knowledge of the disease into the interpretation of data. Moreover, these observations are in agreement with previous studies using the C6 and other ELISA based assays [35, 64, 65]. In a similar study using the C6 test, the State of Virginia showed levels of Lyme seropositive dogs from Southeastern and Mid-Atlantic States to be similar to the levels we found in Texas [66]. These values were significantly lower than the seropositive dogs from other endemic States evaluated in the study, such as Pennsylvania and Maryland. In a more recent study, authors evaluated the sero-prevalence of tick borne pathogens in dogs in North America and the Caribbean using the ELISA SNAP test (IDEXX Inc., Westbrook, ME, USA), and observed that 2.1% of the samples submitted from Texas during years 2008 through 2010 and 2012 were sero-reactive to B. burgdorferi [10]. It is worth mentioning that the difference in percentage might be due to the fact that different testing strategies were used. Hence, our results are in agreement with the fact that Texas is a low prevalence state for LD.

While evaluating the IB results, we observed that the antigens of 23 (OspC), 45, 58 and 66 kDa reacted with sera from both controls and non-control animals, and bands 45 and 58 kDa were more frequently present in control dog samples than non-controls. Therefore, in order to establish robust guidelines for sero-positivity, we agreed to not consider the reaction to these antigens during the test read out. On the other hand, the antigens migrating at 18, 30 and 93 kDa were mostly absent from negative control animals and individuals with low ELISA values, and correlated significantly with the individuals that had high ELISA test values. Animals with reactivity to those antigens, along with other bands, were considered positive for the IB assay. At the same time that we were evaluating the seroprevalence of LD in dogs suspected of a tick borne infection, we obtained a set of dog serum samples seropositive for different Leptospira serovars. We utilized these samples to evaluate possible cross reactivity of the in-house Lyme ELISA test and commercial IB assay. The results obtained suggest that cross-reactivity was not interfering with the detection of seropositive dogs utilizing both the in house ELISA test and the commercially available IB test. Nevertheless, IB positive samples that were Leptospira positive had low Leptospira titers (ranging from 100-800). Therefore, the samples with low Leptospira titers, high Lyme ELISA values and positive IB test are more probable to be Lyme positive than Leptospira positive. In addition, in this study, samples with high Leptospira titers, ranging from 1,600 to12,800 were analyzed and yielded negative Lyme IB test results, supporting the lack of cross-reactivity. This was an interesting finding, since both Leptospirosis and Lyme infections cause renal failure in canids, and as such, in southern states such as Texas a differential diagnosis might be recommended [8, 53, 67-70].

Companion animals have been considered sentinels for LD in endemic and hyperendemic areas [16, 45, 46, 71-75]. Nevertheless, no clear correlation was observed in areas with low incidence of LD. Texas is considered a non-endemic area for this disease with a very low risk for the acquisition of LD in both humans and companion animals. This has been based on calculations utilizing models standardized with data mostly originating in the Northeastern and upper Midwestern US [77-82]. Our study shows that around 10% of the tested animals were sero-reactive by ELISA and immunoblot assay, and this is in agreement with the fact that Texas is a state with very low prevalence of LD. In addition, the majority of seropositive dogs were detected during the months in which we detected the majority of infected ticks (Figure 5). Dogs can be subclinical for months after exposure to infected ticks, and only develop the diseases after encountering environmental stressors. This phenomenon could explain the secondary pick of seroreactive canids observed in this study during the months of May and June, where weather in Texas becomes hot and dry, and could trigger the activation of subclinical infections. In addition, summer weather in Texas is not suitable for Ixodes scapularis tick host-seeking behavior, reducing the probability of infection during those months.  As observed in Figure 4, over 85% of the canine seropositive individuals were distributed in 5 eco-regions of Texas. Interestingly, those eco-regions cover a geographic territory that can provide suitable habitats for the maintenance of I. scapularis ticks, as recently published by Feria-Arroyo and collaborators [54]. These investigators forecasted the potential distribution of the LD vector to 2050, and suggested that even in the context of climate change there will not be major changes in the geographic area that will maintain suitable habitats for I. scapularis in Texas. These observations suggest that the detection of B. burgorferi infected dogs, most importantly those with clinical manifestations, can provide information regarding where this disease could be occurring and the resultant risk for humans. A recent study shows that most of the human reported cases between years 2001 and 2010 co-localize with the detected canine seropositive cases observed in this study, and in particular in the Cross-Timbers eco-region [12]. Consequently, further studies are required utilizing randomized groups of animals to estimate the actual LD sero-prevalence in populations of dogs in Texas, and whether or not they correlate with the reported cases of LD in humans.

In Texas, human LD is diagnosed year around, with no significant peak in any particular season [12]. In our study, the population of dog samples utilized showed an increase in the detection of LD sero-reactive individuals during the months of November through March. This coincides with the time when most of the ticks were submitted for testing by citizens and wildlife management centers to our laboratory (Figure 5A). In addition, the majority of infected ticks were detected in this same time frame (Figure 5B). The ticks analyzed in this study were those submitted by veterinarians and pet owners and therefore we consider them epidemiologically relevant, since these ticks are interacting with the dead end hosts (dogs and humans) and potentially transmitting B. burgdorferi [83-87]. Moreover, serum samples and tick samples were obtained through different sources, which strengthen the results obtained in this study.

Due to the nature of this study, we could not obtain the travel history of each of the animals tested. Thus, we could not account for any visit of these animals to endemic areas in the US. Even though this is a possibility, the time of the year at which most of the cases were detected coincides with the time when most pet owners are participating in outdoor activities within the State of  Texas, such as camping, hunting, and hiking. For instance, white tail deer hunting season lasts from September through February, quail hunting season occurs from October through February, duck hunting season is from October to January, and dove hunting season is also open from September through January, (http://www. animal_listing). These hunting seasons are within the time where most animals were diagnosed, with a tail during spring and early summer that can account for those that could have been infected later in the season, or during visit to endemic areas. A detail to bear in mind is the fact that this study was started in the middle of a historical drought in the US lasting from 2009 till 2012 ( Due to the lack of information on the phenology of I. scapularis in Southern US, our observations suggest that the adult stages of this tick species are mostly active during the fall and winter. No nymphs were received during the study period, so we could not determine whether or not the questing behavior of immature stages was affected by the drought [88-90]. Thus, further studies need to be done in order to determine when the different developmental stages of this tick are active in Southern US, and which stage, nymph or adult, is responsible for the infection of humans and companion animals. These behavioral differences in the tick vector would also explain the low incidence of LD in Southern US, as recently suggested [91, 92]. In these studies, data shows that under southern US conditions of temperature and humidity, I. scapularis nymphs quest at lower heights than under northern conditions [92]. However, both northern and southern I. scapularis populations are very sensitive to temperatures and tend to quest for shorter periods of time at temperatures normally encountered in Southern US [91].

Taken together, our study suggests that sero-reactive domestic canids in Texas were more prevalent during the fall and winter, with a significant reduction of the cases in the spring and summer months. Consequently, we propose that the risk of exposure to LD exists throughout the year in the southern States, with a higher incidence during the cooler months. These observations suggest different seasonality and intensity of exposure in Southern US when compared to the LD transmission cycle in the Northeastern and Midwestern US. An overlap in geographic localization at the eco-region level of I. scapularis tick [54], seropositive dogs and the reported human cases [12], supports the hypothesis that dogs could be sentinel species in areas with low incidence, as it has been shown for parts of the U.S. where LD is endemic or hyperendemic.


Authors want to thank Dr. Mary Nabity for kindly providing the control dog serum samples for the study, and discussions during the preparation of this manuscript, Dr. Adalberto Pérez de León, Dr. Raul F. Medina, Christina Broke and Chloë Snell for their valuable comments to this manuscript. Also, the authors want to thank the Department of Texas Parks and Wildlife for their help providing tick samples from a number of wildlife management areas. Additionally the authors would like to thank Veterinary clinics across the State of Texas, and the Aggieland Humane Society. EMG thanks the Texas A&M University office for Undergraduate Research Scholars for her undergraduate research thesis.

Competing interests:

The authors declare no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Authors’ contributions

EM, AG and MDEG collected and analyzed the samples. SR coordinated the collection samples from TVMDL. MMB conducted all the statistical analysis provided in this study. All authors participated in the design of the study, and in the organization and drafting of this manuscript. All authors read and approved the final manuscript.


This study was funded by Texas AgriLife and the Texas Veterinary Medical Diagnostic Laboratory (TVMDL) seed grant to the project entitled “Improving diagnostic methods for Lyme disease, and epidemiology of human and animal infections in TX” and the Department of Veterinary Pathobiology and Texas A&M University. MDEG has obtained support for this study through the Department of Veterinary Pathobiology, Texas A&M University and AgriLife grant TEXV 6579 (Project I-9524).


The sequences of PCR amplicons are available through GenBank           (NCBI     National                Center    of            Biotechnology Information)        accession              numbers                KJ826413              through KJ826434 for the B. burgdorferi intergenic region (IGR) and KM875668 through KM875675 for the flaB.


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