Ticks used in this study were unfed adult females. As ticks were not fed, modifications of the host to feed ticks were not required, with exception of rabbits. For rabbits, we attached a cotton stockinet on top of the rabbit ear as outlined in the animal use protocol 2011-187 that was approved by Texas A&M University IACUC. The cotton stockinet attachment was used to contain the tick stimulation chamber as detailed below.

Adult Ixodes scapularis and Amblyomma americanum ticks that were used in this study were purchased from the tick rearing facilities at Oklahoma State University (Stillwater, OK, USA). Stimulation of unfed adult A. americanum and I. scapularis ticks to start feeding on different hosts: rabbits, dog, or human was done by exposing ticks to semio-chemicals and temperature as described (Mulenga et al., 2007) with modifications. The modification was that instead of a tick stimulation chamber being made out of a nylon mesh sachet, a tightly capped 2 mL vial that was perforated with a 27-gauge needle was used. The 27-gauge needle perforations were to allow semio-chemicals and body temperature to percolate ticks. Unfed A. americanum (n = 40 in each vial) and I. scapularis (n = 50–80 in each vial) females ticks were enclosed in a stimuli chamber and placed in close proximity with host skin. To ensure the cap did not open for ticks to escape during the stimulation step, the cap was secured with VWR® general-purpose laboratory labeling tape (VWR International, Radnor, PA, USA) and wrapped with Parafilm M® (Bemis Company Inc., Neenah, WI, USA). To expose ticks to semio-chemicals, the stimulation chamber containing ticks was placed in close proximity with host's skin for ~12 h. Three individual hosts: (i) human (Homo sapiens); (ii) New Zealand white rabbit (Oryctolagus cuniculus); and (iii) Dachshund dog (Canis familiaris) were used. To expose the ticks to human semio-chemicals and temperature, the stimuli chamber containing A. americanum or I. scapularis ticks was placed in the front shirt pocket of the volunteer. For exposure to rabbit, stimuli chambers were placed inside cotton stockinet that was attached onto the top of the rabbit ear (Kim et al., 2014). For dogs, stimuli chambers were taped on to the collar/harness. Following exposure to semio-chemicals, ticks were processed for saliva collection as describe below.

I. scapularis ticks mate off the host before interacting with the host (Sonenshine and Roe, 2014). From this perspective, I. scapularis female ticks were pre-mated before being stimulated to start feeding. This was done by putting female and male I. scapularis ticks in a container, and then visually identifying male and female pairs. Male and female I. scapularis pairs were placed in separate container to complete mating. Please note that, since A. americanum ticks mate after taking an initial blood meal (Sonenshine and Roe, 2014), we did not pre-mate females prior to stimulating them to start feeding.

Tick saliva was collected from unfed I. scapularis and A. americanum as previously described (Tirloni et al., 2014; Kim et al., 2016b). For I. scapularis, we collected saliva from unfed non-stimulated (n = 130 ticks) and those that were stimulated to feed on human (n = 80 ticks), rabbit (n = 130 ticks), and dog (n = 80 ticks) hosts. Likewise, for A. americanum, we collected saliva from non-stimulated (n = 40 ticks), and those that were stimulated to feed on human (n = 40 ticks), rabbit (n = 40 ticks), and dog (n = 40 ticks) hosts. Please note that non-stimulated ticks were taken from the incubator (22°C with 90% relative humidity) and were acclimated to room temperature during the saliva collection step. Ticks were rinsed in Milli-Q water, dried on a paper towel and placed dorsal-side down on a glass slide containing tape. Salivation was induced by injecting 0.5–1 μL of 2% pilocarpine hydrochloride (in PBS, pH 7.4) on the ventral side of the lower right coxa using a 34 gauge/0.5 inches/45° angle beveled needle on a model 701 Hamilton syringe (Hamilton Company, Reno, NV, USA). Tick saliva, which in some instances crystalized, was harvested in 2 μL of phosphate buffered saline (PBS) placed on tick mouthparts using a Hamilton syringe every 15 min over ~4 h at room temperature. Saliva protein concentrations were determined using BCA enhanced protocol (BCA Protein Assay, Pierce, Rockford, IL, USA). Tick saliva was lyophilized and stored at −80°C upon use.

Approximately 1 μg of I. scapularis and 1.5 μg of A. americanum total saliva proteins were electrophoresed on 4–20% gradient SDS-PAGE. Gels were silver stained using the Pierce Silver Stain for Mass Spectrometry kit (Thermo Fisher Scientific, Waltham, MA, USA) according to manufacturer's instructions.

Total tick saliva proteins (2 μg, in triplicate) of I. scapularis or A. americanum ticks that were non-stimulated and those that were stimulated to starting feed on different hosts (human, dog, and rabbit) were diluted in 8 M urea/0.1 M Tris, pH 8.5, reduced with 5 mM Tris (2-carboxyethyl) phosphine hydrochloride (TCEP, Sigma-Aldrich, St Louis, MO, USA) and alkylated with 25 mM iodoacetamide (Sigma-Aldrich). Proteins were digested overnight at 37 °C in 2 M urea/0.1 M Tris pH 8.5, 1 mM CaCl₂ with trypsin (Promega, Madison, WI, EUA) with a final ratio of 1:20 (enzyme:substrate). Digestion reactions, in a final concentration of 0.2 μg/mL, were quenched with formic acid (5% final concentration) and centrifuged for debris removal.

Reversed phase pre-columns were prepared in deactivated 250 μm ID/360 μm OD silica capillary (Agilent Technologies, Santa Clara, CA, USA) with a 2 mm Kasil frit at one end. Kasil frits were prepared by dipping 20 cm capillary in 300 μL Kasil 1624 (PQ Corporation, Malvern, PA, USA) and 100 μL formamide solution, curing at 100°C for 3 h and adjusting the length. Pre-columns were packed in-house with 2 cm of 5 μm ODS-AQ C18 (YMC America, INC., Allentown, PA, USA) particles from particle slurries in methanol. Analytical reversed phase columns were fabricated by pulling a 100 μm ID/360 μm OD silica capillary (Molex Polymicro Technologies™, Austin, TX, USA) to a 5 μm ID tip. The same packing material was packed until 20 cm directly behind the pulled tip. Reversed phase precolumns and analytical columns were connected using a zero-dead volume union (IDEX Corp., Upchurch Scientific, Oak Harbor, WA, USA).

Peptide mixtures were analyzed by nanoflow liquid chromatography mass spectrometry using an Easy NanoLC II and a Q Exactive mass spectrometer (Thermo Scientific, Waltham, MA, USA). Peptides eluted from the analytical column were electrosprayed directly into the mass spectrometer. Solutions A and B consisted of 5% acetonitrile/0.1% formic acid and 80% acetonitrile/0.1% formic acid, respectively. The flow rate was set to 400 nL/min. Saliva samples (2 μg per injection) were separated in 155-min chromatographic runs, as follows: 1–10% gradient of solution B in 10 min, 10–40% of solution B in 100 min, 40–50% of solution B in 10 min and 50–90% of solution B in 10 min. Column was held at 90% of solution B for 10 min, reduced to 1% of solution B and re-equilibrated prior to next injection.

The mass spectrometer was operated in a data dependent mode, collecting a full MS scan from 400 to 1,200 m/z at 70,000 resolution and an AGC target of 1 × 10⁶. The 10 most abundant ions per scan were selected for MS/MS at 17,500 resolution and AGC target of 2 × 10⁵ and an underfill ratio of 0.1%. Maximum fill times were 20 and 120 ms for MS and MS/MS scans, respectively, with dynamic exclusion of 15 s. Normalized collision energy was set to 25.

Tandem mass spectra were extracted from Thermo RAW files using RawExtract 1.9.9.2 (McDonald et al., 2004) and searched with ProLuCID (Xu et al., 2015) against a non-redundant tick databases. I. scapularis peptides were searched against the database containing an Ixodidae sequences from NCBI (62,246 sequences and reverse sequences). For A. americanum, we searched peptides against an in house database of translated whole tick and dissected organ transcriptome (BioProject accession number PRJNA226980). Searches were done using Integrated Proteomics Pipeline—IP2 (Integrated Proteomics Applications, Inc.). The search space included all fully-tryptic and half-tryptic peptide candidates. Carbamidomethylation on cysteine was used as static modification. Data was searched with 50 ppm precursor ion tolerance and 20 ppm fragment ion tolerance.

The validity of the peptide spectrum matches (PSMs) generated by ProLuCID was assessed using Search Engine Processor (SEPro) module from PatternLab for Proteomics platform (Carvalho et al., 2015). Identifications were grouped by charge state and tryptic status, resulting in four distinct subgroups. For each group, ProLuCID XCorr, DeltaCN, DeltaMass, ZScore, number of peaks matched and secondary rank values were used to generate a Bayesian discriminating function. A cut-off score was established to accept a false discovery rate (FDR) of 1% based on the number of decoys. This procedure was independently performed on each data subset, resulting in a false-positive rate that was independent of tryptic status or charge state. Additionally, a minimum sequence length of six residues per peptide was required. Results were post-processed to only accept PSMs with <10 ppm precursor mass error. A Principal Component Analysis (PCA) plot, performed using PatternLab's Buzios module (Carvalho et al., 2015), was employed to aid in interpreting similarities among samples. Venn's four-set diagrams were generated from the output of PatternLab's Birds Eye view report. Proteins were grouped by maximum parsimony and the presence of proteins in at least two out of three replicates was required for each condition.

Volcano plots were generated by a pairwise comparison between non-stimulated and stimulated tick saliva using PatternLab's TFold module, which uses a theoretical FDR estimator to maximize identifications satisfying both a fold-change cut-off that varies with the t-test p-value as a power law and a stringency criterion that aims to fish out proteins of low abundance that are likely to have had their quantitation compromised (Carvalho et al., 2015). The following parameters were used to select differentially expressed proteins: proteins were grouped by maximum parsimony, spectral count data was normalized using normalized spectral abundance factor (NSAF) (Zybailov et al., 2006), two (out of the three runs) non-zero replicate values were required for each condition, and a BH q-value was set at 0.02 (2% FDR). Low abundant proteins were removed using an L-stringency value of 0.2.

To get insight on the nature of the identified protein sequences, BLASTp searches against several databases were performed. To functionally classify the protein sequences, a program written and provided by Dr. José M. C. Ribeiro in Visual Basic 6.0 (Microsoft, Redmond, WA, USA) was used (Karim et al., 2011). The functionally annotated catalog for each dataset was manually curated and plotted in a hyperlinked Excel spreadsheet designed as Table S1 (for I. scapularis) and Table S2 (for A. americanum).

We have recently identified proteins in saliva of I. scapularis ticks that were fed every 24 h on rabbits (Kim et al., 2016b). To determine if some proteins reported here were injected into the host during tick feeding, unfed I. scapularis tick saliva proteome from this study were scanned against published I. scapularis proteome.

Proteomic profiles were compared across samples as functional classes or individual proteins. To determine the relative abundance of proteins, normalized spectral abundance factor (NSAF) was used in a label-free relative quantification approach (Paoletti et al., 2006). Mean NSAF values from the two or three replicates were determined and combined according to functional class, and then divided by the total NSAF for the respective sample. NSAF as an index for relative protein abundance was input in Microsoft Excel (Microsoft, Redmond, WA, USA) as percentage of the total NSAF for respective samples, and visualized on pie charts according to protein classes. To visualize relative expression patterns on a heat map, NSAF values were normalized using Z-score. Normalized NSAF values were used to generate heat maps using the heatmap.2 function from the gplots package in R.

The mass spectrometry proteomics raw data have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD00712.

The hard tick's adaptation to feed on different animal hosts is central to TBD epidemiology as it facilitates the tick's movement of TBD agents from their wild animal reservoir hosts to humans, farm and companion animals. Despite its centrality, the molecular basis of how the tick adapts to feed on different hosts has not been fully evaluated. The tick feeding style of lacerating host tissue and then sucking up host blood that bleeds into the wounded area (commonly known as the tick-feeding site) is thought to stimulate host tissue repair responses that are aimed at stopping further blood loss. However, ticks ensure a full blood meal by secreting a cocktail of proteins that disarm the host's tissue repair response (Francischetti et al., 2009). There is also evidence the tick may express specific sets of genes in preparation to start feeding (Mulenga et al., 2007; Lew-Tabor et al., 2010; Rodriguez-Valle et al., 2010). This study was initiated to answer the question of whether or not the molecular preparation to start feeding by ticks such as I. scapularis and A. americanum that are adapted to feed on a wide range of hosts utilized was the same regardless of the host. Findings in this study suggest that the tick's molecular preparation to start feeding could be host-specific as indicated by SDS-PAGE and silver staining analysis that revealed unique protein profiles in saliva of I. scapularis and A. americanum ticks that were stimulated to start feeding on rabbits (SR), dog (SD), human (SH), and those that were not-stimulated (NS) (Figure 1). Solid (SB) and broken (BB) line boxes respectively highlight similarity and differences of detectable protein band patterns in saliva of ticks that were stimulated to feed on different hosts (Figure 1). Whereas we observed similarities (solid boxes) and differences (broken boxes) in protein banding patterns in saliva of I. scapularis ticks that were stimulated to feed on dogs and humans (Figure 1A), there are no apparent similarities among protein banding patterns in saliva of the four A. americanum treatments (Figure 1B). A notable observation in Figure 1B is that protein banding patterns in saliva of dog and rabbit-stimulated A. americanum ticks were closely similar, while those exposed to humans show more differences. Due to insufficient sample amounts, non-stimulated and rabbit-exposed I. scapularis tick saliva proteins were not analyzed by SDS-PAGE (Figure 1A). From this perspective, our SDS-PAGE analysis in Figure 1A was limited, and thus its unclear if observations in A. americanum were consistent with those in I. scapularis. The observation that the protein profile in saliva of non-stimulated A. americanum ticks is different from those that were stimulated to start feeding (Figure 1B) further demonstrates that stimulating ticks to start feeding impacted proteins that were secreted into tick saliva.

There is evidence that exposing the animal to high temperature stress can affect expression levels of different proteins (Villar et al., 2010). However, given that the respective axillary temperatures of dogs (~38°C), human (~37°C), and rabbits (~36°C) are not dramatically different (Vadlejch et al., 2010; Goic et al., 2014), it is unlikely that the observed differences in protein profiles were temperature dependent. For our non-stimulated tick cohorts, saliva was collected from ticks that were obtained straight from the incubator at 22°C and acclimated to room temperature, which is normally around 25°C. Thus, observed shift in protein profiles between NS and those that were stimulated to start feeding on different hosts (which have higher respective axillary temperatures) cannot be ruled out. On the other hand, studies have been shown that ticks respond to mechano-, chemo-, and thermo-sensation and are able to induce different electrophysiological responses (Soares and Borges, 2012; Sonenshine and Roe, 2014). Upon contact with host, release of host-derived semio-chemicals and their interaction with tick sensory organs may result in different electrophysiological responses leading secretion of different proteins during host stimulation process. Since ticks used in this study did not come into contact with the host's skin, tick saliva proteins described reported may not include those are responsive to mechano-sensory stimulation.

To get further insights into differences observed among saliva proteomes from ticks that were exposed to different hosts, we proceeded to identify proteins by LC-MS/MS using an in-solution digestion approach as described (Tirloni et al., 2014; Kim et al., 2016b). We respectively identified a total of 276 and 340 non-redundant I. scapularis (Figure 2A) and A. americanum (Figure 2B) tick saliva proteins that were determined authentic as they were detected in two or all of the three LC-MS/MS runs that were done for each sample (Figure 2, Tables S1, S2). The remaining 69 (I. scapularis) and 57 (A. americanum) proteins that were detected in only one of the three runs were considered low confidence hits and not further discussed (Tables S1, S2). Figure 2 summarizes the overall total proteins that were identified in saliva of ticks not stimulated to start feeding (NS) and those stimulated to start feeding on human (SH), dog (SD), and rabbit (SR). Of the 276 unique I. scapularis tick saliva proteins (Table S1), 66, 189, 186, and 165 were respectively identified in saliva of NS, SH, SD, and SR ticks (Figure 2A). Of these, 55 were common to all treatments, while 2, 35, 35, and 34 proteins were respectively unique to NS, SH, SD, and SR ticks (Figure 2A). Likewise, in A. americanum we respectively found 245, 192, 288, and 93 proteins in saliva of NS, SH, SD, and SR ticks (Figure 2B). Of these, 67 were common to all treatments, while 19, 12, 59, and 9 proteins were respectively unique to NS, SH, SD, and SR ticks (Figure 2B). Functional annotations classified both I. scapularis and A. americanum tick saliva proteins into 28 protein classes (Figure 3, Tables S1, S2). Based on the total sum of normalized spectral abundance factor (NSAF) (Figure 3), the predominant classes of proteins in this study include secreted conserved proteins (unknown functions), proteinase inhibitors, lipocalins, extracellular matrix/cell adhesion, heme/iron metabolism, signal transduction and immunity-related proteins (Figure 3).

Graphic visualization data in Figure 4 summarizes the Z-score of normalized NSAF values for each of the 28 functional classes identified in I. scapularis (Figure 4A) and A. americanum (Figure 4B). These data reveal two general trends: (i) the tick might inject the same protein at different levels into different hosts, and that (ii) protein composition in saliva of different tick species that feed on the same host is likely different (Figure 4). For instance, saliva of SR I. scapularis ticks had high abundance of heme/iron metabolism (22.4%) followed by extracellular matrix/cell adhesion (7%), oxidant metabolism/detoxification (6.2%), cytoskeletal (4.6%), metabolism (amino acid, carbohydrate and energy) (2%), proteasome machinery (1.1%), nuclear regulation (0.8%), conserved protein with unknown functions (0.6%), protein modification (0.4%), protein synthesis machinery proteins (0.4%), and transport/storage (0.02%) (Figure 4A and Table S1). In contrast, saliva of SR A. americanum ticks had high abundance of extracellular matrix/cell adhesion (18.6%) and proteinase inhibitors (18.2%), followed by heme/iron metabolism (11.1%), immunity-related (4.8%), metabolism of energy proteins (1.4%), cytoskeletal (0.7%), protein synthesis machinery (0.5%), and proteasome machinery (0.1%) (Figure 4B and Table S2).

An interesting finding in our data is that the 55 and 67 proteins that were respectively found in all I. scapularis and A. americanum treatments (Figures 2A,B) belonged to the same functional classes (Figure S1, Tables S1, S2). These data suggest that A. americanum and I. scapularis utilized a core set of functionally similar proteins that regulated key host defense pathways to successfully feed. Although functional role(s) of proteins in Figure S1 remain to be determined, available evidence indicate that some of these proteins regulated important tick feeding pathways. For instance, a cross-tick species conserved AV422 protein that was originally identified among genes that were up regulated in A. americanum ticks that were stimulated to feed on cattle (Mulenga et al., 2007) and was injected into animals during A. americanum (Mulenga et al., 2013), I. scapularis (Kim et al., 2016b), R. microplus (Tirloni et al., 2014), and Haemaphysalis longicornis (Tirloni et al., 2015) feeding is an inhibitor of blood clotting and platelet aggregation (Mulenga et al., 2013). Likewise, EEC19556.1, which was found in all I. scapularis treatments (Table S1) is 99% identical to a serine protease inhibitor (serpin, AID54718.1) anti-coagulant and inhibitor of thrombin (Ibelli et al., 2014) that is injected into rabbits during I. scapularis feeding (Kim et al., 2016b).

In order to investigate if shared proteins were differentially secreted I. scapularis and A. americanum when stimulated to start feeding on different hosts, pairwise comparison analyses using the PatternLab's TFold module (Carvalho et al., 2015) were conducted (Figure 5, Tables S4, S5). This analysis demonstrated that some of the shared proteins were secreted at equivalent levels (red dots), not significantly different (green and yellow dots), and significantly at different levels (blue dots) (Figure 5). Based on fold change (FC), differences in abundance ranged between 18.0 and 1.2 for I. scapularis (Table S4) and between 40 and 1.2 for A. americanum (Table S5). Consistent with our analysis in Figure 4, majority of SR I. scapularis tick saliva proteins were secreted at high concentrations when compared to either SD or SH. When compared to SD or SH, some of the most abundant proteins in saliva of SR I. scapularis ticks, included superoxide dismutase (EEC10196.1, FC: 17.7, and FC: 9.3), a glyceraldehyde-3-phosphate dehydrogenase—GAPDH (JAA68969.1, FC: 10.7), a tropomyosin (JAB83342.1, FC: 9.9), a thymosin (JAA70823.1, FC: 7.3), a fructose 1,6-biphosphate aldolase (EEC14101.1, FC: 8.3), and a creatine kinase (JAB78095.1, FC: 6.9). When paired with either rabbit or human, proteins with higher FC in saliva of SD I. scapularis ticks, include oxidase/peroxidase enzyme (EEC07462.1, FC: 18.0), a serine protease (EEC02857.1, FC: 6.7), a secreted salivary gland peptide (EEC14213.1, FC: 6.7), a peroxinectin (EEC08358.1, FC: 6.5), a microplusin (AAY66495.1, FC: 4.8), a GAPDH (JAA68969.1, FC: 4.7), and a 14-3-3 zeta protein (JAB76832.1, FC: 4.3). Likewise, for human-exposed ticks an oxidase/peroxidase (EEC07462.1, FC: 9.6), a secreted salivary gland peptide (EEC14213.1, FC: 7.5), an insulin growth factor-binding protein (EEC07853.1, FC: 5.0), a metalloprotease (AAM93652.1, FC: 7.0), and a secreted protein (EEC14470.1, FC: 4.0) were identified with higher FC (Table S4).

Although similar to I. scapularis (Table S4), proteins in saliva of A. americanum were likely secreted at high concentration when stimulated to feed on rabbits than either dog or human (Table S5). It is notable that fewer proteins were differentially abundant in SH (n = 2) or SD (n = 4) A. americanum tick saliva when compared with SR saliva (Figure 5B and Table S5). Please note that four of the six proteins that were differentially abundant in SH compared to SR A. americanum saliva (Figure 5B) are potentially contaminating keratins (Table S5), and hence the six blue dots in Figure 5. Whether or not the observation here is a consistent biological phenomenon remains to be determined. We also think that this observation could be explained by the limitation of our study: we searched A. americanum peptides against a database of a translated A. americanum transcriptome that was generated from ticks that were fed on rabbits. We anticipate that if the A. americanum genome became available, additional host-dependent A. americanum tick saliva protein secretions will be identified.

Despite the fact that we identified a limited number of differentially abundant A. americanum proteins, the secretion dynamics of AV422 (Mulenga et al., 2007) was similar in both A. americanum and I. scapularis (Tables S4, S5). Our data here show this protein is secreted at high abundance when A. americanum is stimulated to feed on humans (FC: 1.7) and dogs (FC: 2.2). Similarly, in I. scapularis, AV422 homolog is secreted at high abundance when I. scapularis is stimulated to feed on humans (FC: 1.6) and dogs (FC: 1.8). It will be interesting to further investigate the role(s) of I. scapularis AV422 homolog in tick feeding were similar to those observed A. americanum (Mulenga et al., 2013).

The observation that, A. americanum ticks may secrete fatty acid-binding protein at FC of 40.2 (Aam-134, Table S5) represents the most abundantly secreted protein in saliva of SR A. americanum tick saliva. This protein was identified exclusively in saliva of SR and SD ticks and appears more abundant in the former (Table S5). Fatty acid-binding proteins have been identified in helminth secretions (Morphew et al., 2007) and in tick saliva. This protein has been described to modulate human monocyte-derived macrophages and generate M2 macrophages (MΦ) activated by the alternative pathway (Figueroa-Santiago and Espino, 2014). M2 MΦ are characterized by secretion of anti-inflammatory cytokines (Gordon, 2003). Since ticks have to evade the host's inflammation defense, could the fatty acid-binding protein being among tick proteins that modulate host defense? It would be interesting to address if the A. americanum fatty acid-binding protein has similar effects on host immune modulation. Other proteins showing high FC values in saliva of A. americanum SR ticks are glycine-rich proteins: Aam-178421 (FC: 29.1), Aam-1227 (FC: 29.0), Aam-327 (FC: 13.1) when pairing with SH; and Aam-179267 (FC: 18.6), Aam-177922 (FC: 14.2), and Aam178421 (FC: 12.8) when pairing with SD (Table S5). Glycine-rich proteins are extracellular matrix proteins and/or structural proteins thought to play an important role in attachment to the host, as they are present in cement material secreted by salivary glands during feeding process (Bishop et al., 2002; Maruyama et al., 2010). Since these proteins are secreted in the early stage of tick feeding, we suggest that glycine-rich proteins could be associated with tick cement formation, securely anchoring ticks onto host skin during the prolonged tick-feeding period. Another set of proteins with high FC values include a hemelipoprotein (FC: 21.3), a GAPDH (FC: 16.0), several serpins (FC: 15.9 to 1.5), among others (Table S5). More details about the computational comparison are available in Figure 5, Tables S4, S5.

The observation of apparent differential expression is not likely peculiar to this study. In a lone study, tick proteins involved in blood digestion and reproduction were overrepresented in R. microplus ticks that fed on cattle when compared to ticks that fed on white-tailed deer (Popara et al., 2013). In a related study, I. scapularis displayed variable fibrinogenolytic activities upon feeding on mice with different immune backgrounds (Vora et al., 2017).

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.