The Unfolded-Protein Response Triggers the Arthropod Immune Deficiency Pathway

ABSTRACT The insect immune deficiency (IMD) pathway is a defense mechanism that senses and responds to Gram-negative bacteria. Ticks lack genes encoding upstream components that initiate the IMD pathway. Despite this deficiency, core signaling molecules are present and functionally restrict tick-borne pathogens. The molecular events preceding activation remain undefined. Here, we show that the unfolded-protein response (UPR) initiates the IMD network. The endoplasmic reticulum (ER) stress receptor IRE1α is phosphorylated in response to tick-borne bacteria but does not splice the mRNA encoding XBP1. Instead, through protein modeling and reciprocal pulldowns, we show that Ixodes IRE1α complexes with TRAF2. Disrupting IRE1α-TRAF2 signaling blocks IMD pathway activation and diminishes the production of reactive oxygen species. Through in vitro, in vivo, and ex vivo techniques, we demonstrate that the UPR-IMD pathway circuitry limits the Lyme disease-causing spirochete Borrelia burgdorferi and the rickettsial agents Anaplasma phagocytophilum and A. marginale (anaplasmosis). Altogether, our study uncovers a novel linkage between the UPR and the IMD pathway in arthropods.

To evaluate whether tick-borne pathogens induce the UPR in I. scapularis, we quantified UPR-associated gene expression in A. phagocytophilum-infected nymphs. Specifically, we evaluated expression levels of BiP, the three UPR receptors (IRE1a, PERK, and ATF6), and molecules associated with the IRE1a pathway, XBP1 and TRAF2. Relative to uninfected ticks (Fig. 1B, dotted baseline), significant increases were observed with BiP, ire1a, and traf2, suggesting that the tick UPR responds to infection (Fig. 1B).
To determine how the UPR impacts pathogen survival in ticks, we used pharmacological inducers or RNAi with the ISE6 I. scapularis cell line. Tick cells were treated with low doses of either thapsigargin or tunicamycin to induce ER stress prior to A. phagocytophilum infection. Thapsigargin inhibits the sarco/endoplasmic reticulum Ca 21 ATPase (SERCA), which decreases calcium levels in the ER (60). Tunicamycin blocks N-linked glycosylation, leading to an increase of misfolded proteins (61). Both treatments resulted in significantly less A. phagocytophilum ( Fig. 1C and D). We also used an RNAi-based approach to overactivate the UPR by decreasing expression of the negative regulator BiP. In agreement with pharmacological induction, transcriptional silencing of BiP caused a decrease in A. phagocytophilum colonization (Fig. 1E). Altogether, this demonstrates that A. phagocytophilum induces the UPR in ticks, which functionally restricts bacterial colonization and survival.
Infection induces IRE1a activation, but not XBP1. Transcripts induced by A. phagocytophilum are associated with the IRE1a signaling axis ( Fig. 1A and B), which is the most conserved branch of the UPR among eukaryotes (62). When activated, IRE1a autophosphorylates and either splices the mRNA xbp1 (X-box binding protein 1) or signals through TRAF2 (36, 37, 46, 57) (Fig. 1A). Unspliced xbp1 mRNA (xbp1 U ) is held in an inactive state in the cytoplasm by forming a hairpin structure that inhibits translation. The RNase domain of IRE1a splices an internal intron from xpb1 U , allowing it to be translated into a protein that functions as a transcription factor (57,(63)(64)(65)(66) (Fig. 1A). Alternatively, IRE1a can recruit the signaling molecule TRAF2 to produce proinflammatory responses through NF-k B signaling (36)(37)(38)46). We aligned mammalian sequences from the IRE1a pathway with tick homologs and observed sequence similarity with BiP, IRE1a, XBP1, and TRAF2 ( Fig. S1A to D). Notably, the IRE1a kinase domain, RNase domain, and the activity-inducing phosphoserine (Fig. S1B) were well conserved with human sequences. Given this sequence conservation, we used an antibody against human phosphorylated IRE1a to examine the posttranslational activation status of IRE1a in ticks. Upon treatment with the UPR inducers thapsigargin and tunicamycin, increased IRE1a phosphorylation was observed in ISE6 tick cells by immunoblotting, as expected (Fig. S2A). A. phagocytophilum and B. burgdorferi also induced IRE1a phosphorylation in ISE6 cells, indicating that infection induces receptor activation ( Fig. 2A  and B). A small-molecule inhibitor, KIRA6 (67), successfully blocked IRE1a phosphorylation during infection ( Fig. 2A and B). Inhibiting IRE1a phosphorylation caused significantly increased infection in tick cells (Fig. 2C). Similarly, knocking down the expression of ire1a through RNAi also increased A. phagocytophilum bacterial burden ( Fig. 2D;  Fig. S2B). These data show that IRE1a signaling in ticks is activated by infection and restricts bacterial colonization in vitro.
To delineate the signaling events downstream from IRE1a, xbp1 U was next examined in infected ISE6 cells. Primers flanking the xbp1 intron (Fig. S1E) were used to differentiate spliced and unspliced transcripts by PCR. Unspliced xbp1 U migrates as a single 459-bp band. In contrast, spliced xbp1 S presents as a trimer on an agarose gel, consisting of spliced transcripts (xbp1 S , 434 bp), unspliced transcripts (xbp1 U ), and an xbp1 U -xbp1 S heterodimer that is an artifact of PCR and migrates slightly higher. Spliced xbp1 S was observed in thapsigargin-treated tick cells under all conditions. In contrast, neither A. phagocytophilum or B. burgdorferi induced xbp1 U splicing at any time points in vitro ( Fig. 2E and F). We next probed in vivo samples from replete I. scapularis nymphs that were fed on either uninfected mice or mice infected with A. phagocytophilum or B. burgdorferi. Across all samples, xbp1 U remained unspliced (Fig. 2G). These results indicate that although the tick IRE1a is activated by infection and restricts bacterial burden, this phenotype is not carried out through XBP1 activity.
Since XBP1 is not responsive to infection, we sought to determine if TRAF2 has a functional role during pathogen colonization. Reducing the expression of traf2 through RNAi in Ixodes ISE6 cells caused a significant increase in A. phagocytophilum (Fig. 2H), correlating with the phenotype observed when ire1a transcripts were silenced (Fig. 2D). These data, together with upregulated traf2 expression in A. phagocytophilum-infected I. scapularis nymphs (Fig. 1B), led us to further interrogate whether IRE1a signals through TRAF2 to restrict pathogen colonization.
IRE1a interfaces with TRAF2 in I. scapularis ticks. Aligning sequences from humans and ticks reveals that the Ixodes TRAF2 is fundamentally unique compared to the mammalian homolog (Fig. S3A). The Ixodes TRAF2 lacks a RING (really interesting new gene) domain that is necessary for ubiquitin ligase activity (68). The Ixodes TRAF2 also has a reduced TRAF-N domain, which is responsible for bridging interactions with other proteins (68). Given these differences, we performed homology modeling and a prediction-driven docking approach (69) with the I. scapularis IRE1a and TRAF2 proteins to gain insight into how they interact. BLAST was used to identify the human TRAF2 crystal structure (68) (PDB code 1CA9) as a modeling template for Ixodes TRAF2. The modeled form of the Ixodes TRAF2 C-terminal region features part of a coiled-coil domain and the highly conserved TRAF-C domain (Fig. S3B). In addition, the homology model is a trimer where the coiled-coil domain is a single alpha helix and the TRAF-C domain forms an eight-stranded antiparallel b-sandwich. Next, the human IRE1a crystal structure (72) (PDB code 6URC) was identified by BLAST as a homology template for modeling the cytosolic RNase/kinase domain of I. scapularis IRE1a. The structure was modeled in the active-state quaternary structure proposed to be necessary for autophosphorylation and RNase activity (73) (Fig. S3C and D).
We then modeled the Ixodes IRE1a-TRAF2 complex using a prediction-driven docking approach (69). This tactic combines the utility of interface prediction with ab initio docking and is a useful alternative to ab initio docking alone when protein-protein complex formation is being examined. CPORT (consensus prediction of interface residues in transient complexes) (69) was used to assign active and passive residues at the interface of the trimeric TRAF-C domains and the RNase/kinase domain of IRE1a (Fig. 3A). Residues were then used to filter the docking process by HADDOCK 2.2 (74), which optimizes residue conformations at the interface before proceeding to refinement. The docking model places the trimeric TRAF2 interface at the kinase domain of IRE1a with a buried surface area of 3,262.16 Å 2 (Fig. 3B). Importantly, trimeric TRAF2 is positioned in a manner that does not interfere with the IRE1a dimer interface and is away from the C-terminal transmembrane domain (Fig. 3B, circled) that anchors IRE1a to the ER (Fig. 3B). Five salt bridge interactions were identified that define how the TRAF2 trimer is positioned onto the kinase domain of IRE1a (Fig. 3C). Each chain of TRAF2 participates in salt bridge interactions with the kinase domain of IRE1a. Therefore, the oligomeric state of TRAF2 seems to play an important role in docking specificity with the RNase/kinase domain of IRE1a. Altogether, in silico docking analyses with Ixodes IRE1a and TRAF2 suggest that these two molecules can directly interface with one another.
To experimentally validate that IRE1a and TRAF2 specifically interact, we used a human embryonic kidney (HEK) 293T cell transfection system with plasmids expressing Ixodes IRE1a and TRAF2 fused to affinity tags (Fig. 3D). Recombinant protein expression was confirmed by immunoblotting transfected cells with antibodies for FLAG and hemagglutinin (HA) tags (IRE1a-FLAG and TRAF2-HA). When Ixodes IRE1a and TRAF2 are coexpressed, immunoprecipitation with antibodies against the FLAG tag demonstrates that IRE1a specifically pulls down TRAF2 and vice versa (Fig. 3D). Altogether, these data demonstrate that Ixodes IRE1a and TRAF2 directly and specifically interact.
Ixodes IRE1a and TRAF2 restrict in vivo bacterial colonization in ticks. We next determined whether the pathogen-restricting activity of Ixodes IRE1a and TRAF2 observed in vitro had similar impacts in vivo. To knock down gene expression, unfed I. scapularis nymphs were microinjected with small interfering RNA (siRNA) targeting ire1a and traf2 or with a scrambled control RNA (scRNA). Nymphs were rested overnight and then fed to repletion on A. phagocytophilum-infected mice. Gene silencing and bacterial burden were both quantified by quantitative reverse transcriptase PCR (qRT-PCR). Similar to in vitro experiments, reducing the expression of ire1a and traf2 led to an increase in A. phagocytophilum burdens in I. scapularis nymphs ( Fig. 4A and B).
I. scapularis take a blood meal once per life stage, with ticks initially becoming infected during the larval phase (75). Since gene expression can vary depending on arthropod life stage (76)(77)(78), we examined the impact of IRE1a and TRAF2 on pathogen colonization in larvae. We silenced ire1a and traf2 in I. scapularis larvae using a modified immersion protocol where ticks were submerged in siRNA or scrambled controls overnight (79). Following immersion, larvae were rested for 24 h before feeding to repletion on A. phagocytophilum-infected mice. Significant knockdown of ire1a and traf2 was observed in siRNA-treated larvae with this method, which caused an increase in A. phagocytophilum numbers ( Fig. 4C and D).

UPR-Initiated IMD Pathway mBio
Soon after A. phagocytophilum is acquired, the bacteria migrate to the salivary glands, where they persist throughout the tick life cycle (75,80,81). To understand how IRE1a influences bacterial colonization in tick tissue subsets, we employed an ex vivo tick organ culture system (82,83). Midguts and salivary glands from adult I. scapularis ticks were dissected and treated with the IRE1a inhibitor KIRA6 prior to infection with A. phagocytophilum (Fig. 4E). Similar to in vitro and in vivo findings, inhibiting the activity of IRE1a led to significantly higher A. phagocytophilum burdens in ex vivo salivary gland and midgut cultures ( Fig. 4F and G), demonstrating that this signaling axis functionally restricts bacterial colonization in disparate tick tissues.
We next asked whether the activity of IRE1a-TRAF2 signaling was restrictive to different tick-borne microbes, such as the Lyme disease-causing spirochete B. burgdorferi. Expression of ire1a and traf2 was knocked down through RNAi in both I. scapularis nymphs and larvae using the methods described above, and ticks were fed to repletion on B. burgdorferiinfected mice. In agreement with the phenotype observed with A. phagocytophilum, significantly higher B. burgdorferi levels were observed in siRNA-treated ticks at both the nymph ( Fig. 5A and B) and larval ( Fig. 5C and D) stages. These data show that IRE1a-TRAF2 signaling is broadly responsive to multiple I. scapularis-transmitted pathogens and is functionally restrictive to microbial colonization during different tick life stages.
The IMD pathway is triggered by IRE1a. TRAF2 is a component of the mammalian TNFR network, which is functionally analogous to the arthropod IMD pathway. This parallel led us to ask whether the antimicrobial activity of the Ixodes IRE1a-TRAF2 axis operates through arthropod immunity. AMPs specific to the IMD pathway have not yet been identified in ticks. Instead, the Drosophila S2* cell line can be used as a surrogate model to quantify pathway-specific AMPs (32). To examine whether ER stress induces an immune response in the absence of microbes, we treated S2* cells with the UPR inducer thapsigargin. AMPs corresponding to the IMD pathway (diptericin, attacin A, and cecropin A2) (84) were significantly induced in a dose-dependent manner compared to unstimulated controls (Fig. S4A). In contrast, the Toll pathway AMP IM1 (84-86) was not significantly different, demonstrating that ER stress leads to IMD pathway activation independent of microbial agonists.
It is known that the IMD pathway is responsive to the tick-transmitted pathogens A. phagocytophilum and B. burgdorferi (32,33). Since tick-borne microbes also activate the UPR ( Fig. 1B and 2A) and ER stress induces the IMD network (Fig. S4A), we asked whether blocking IRE1a during infection would inhibit the IMD pathway. S2* cells that were treated with the IRE1a inhibitor KIRA6 prior to A. phagocytophilum or B. burgdorferi infection showed significantly reduced IMD pathway AMPs ( Fig. S4B and C).
We next examined whether the tick IMD pathway underwent a similar UPR-driven activation event. Relish is the transcription factor associated with IMD pathway activation. Similar to what was observed in Drosophila S2* cells, ISE6 cells that were treated with UPR stimulator thapsigargin or tunicamycin showed an increase in Relish activation (Fig. 6A). We next asked if inhibiting IRE1a would block activation of the IMD pathway in ticks. ISE6 cells were stimulated with A. phagocytophilum and B. burgdorferi alone or were pretreated with the IRE1a inhibitor KIRA6 before infection. Pretreatment with KIRA6 resulted in a decline in Relish activation ( Fig. 6B and C), indicating that infection-induced IMD pathway activation occurs through IRE1a. Collectively, our results provide strong evidence that the IRE1a-TRAF2 axis functions as an IMD pathway-activating mechanism.
Ixodes IRE1a-TRAF2 signaling potentiates reactive oxygen species. An immune mechanism complementary to the IMD pathway is the production of reactive oxygen species (ROS), which cause bactericidal damage to nucleic acids, proteins, and membrane lipids (16,87,88). Because B. burgdorferi and A. phagocytophilum are both sensitive to killing by ROS (89)(90)(91)(92) and the mammalian UPR can lead to ROS production (53, 93), we investigated whether ROS can be induced by the Ixodes IRE1a-TRAF2 pathway. ISE6 cells were stimulated with either thapsigargin, tunicamycin, or a vehicle control and monitored for ROS with the fluorescent indicator 29,79-dichlorofluorescein diacetate. Pharmacological inducers caused significantly higher fluorescence, indicating that the tick UPR potentiates ROS (Fig. 6D). Infection with A. phagocytophilum and B. burgdorferi also elicited ROS production in tick cells ( Fig. 6E and F). Pretreating ISE6 cells with the ROS-inhibiting agent diphenyleneidonium chloride (DPI) prior to infection reduced fluorescence, as expected. Importantly, blocking IRE1a activity with KIRA6 either reduced or completely mitigated ROS ( Fig. 6E and F), demonstrating that infection-induced ROS production is potentiated by IRE1a. IRE1a-TRAF2 signaling restricts pathogens across tick vectors. Since the UPR is conserved across eukaryotes, we explored the possibility that the microbe-restricting activity of IRE1a-TRAF2 signaling could functionally impact other arthropod vectors. D. andersoni ticks are important disease vectors that transmit several pathogens, including the obligately intracellular rickettsia A. marginale (94). When inducing the UPR in the D. andersoni tick cell line DAE100 with tunicamycin and thapsigargin ( Fig. 7A and B) or blocking IRE1a with KIRA6 (Fig. 7C), we observed significant changes in A. marginale invasion and replication, comparable to what was observed with I. scapularis and A. phagocytophilum (Fig. 1C and D and 2C). Moreover, higher bacterial loads were also observed in D. andersoni ex vivo midgut and salivary gland cultures when IRE1a activity was blocked with KIRA6 ( Fig. 7D to F). Altogether, this demonstrates that the microbe-restricting activity of IRE1a-TRAF2 signaling is conserved across tick species

DISCUSSION
How arthropod immunity responds to infection is a fundamental factor influencing the ability of vectors to harbor and transmit pathogens (2)(3)(4)(5)(6)(7)(8). The IMD pathway is increasingly recognized as being divergent across species, with classically defined upstream regulators missing in many arthropod genomes (16-29, 31, 34). This suggests that an alternative activation mechanism exists. In this article, we demonstrate that the I. scapularis IMD pathway is initiated through the IRE1a-TRAF2 axis of the UPR. Colonization and replication of A. phagocytophilum and B. burgdorferi are restricted in ticks by Ixodes IRE1a and TRAF2 both in vitro and in vivo. Moreover, we show that IMD pathway activation and ROS production in response to A. phagocytophilum and B. burgdorferi are dependent on IRE1a activity and that this mode of antibacterial restriction is conserved across arthropods. Collectively, our findings provide an explanation for how the core IMD pathway is activated in the absence of canonical upstream regulators.
To our knowledge, this is the first time that cellular stress responses have been implicated in influencing vector competency. Why host cell stress responses are triggered by A. phagocytophilum and B. burgdorferi remains unclear. Ticks do not appear to suffer pathological consequences from the microbes they transmit. The connection between host cell stress and immune outcomes supports a model where transmissible pathogens would benefit most by decreasing infection-induced stress. This model is reenforced by the absence of common inflammatory PAMPs in many tick-transmitted pathogens. For example, all Ixodes-transmitted bacteria lack lipopolysaccharide (LPS) and DAP -PGN (95-98). B. burgdorferi flagella are housed in the periplasm, effectively shielded from recognition by host cells (99). During coevolution with ticks, Ixodes- Nevertheless, our data show that A. phagocytophilum and B. burgdorferi exert at least some stress on ticks. Since immune responses are energetically costly to the host (100, 102), we speculate that the tick response is tuned to match the level of threat imposed by infection, ultimately striking a balance that conserves resources and preserves tick fitness (9, 101).
Our findings indicate a mechanism of IMD pathway activation that deviates from the classically defined paradigm where pattern recognition receptors (PRRs) sense bacterially derived PAMPs. Both intracellular and extracellular pathogens exert stress on the host, which can be caused by secreted toxic by-products, competition for nutrients, and/or physical damage to host cells/organ systems (50). For example, B. burgdorferi is an extracellular spirochete and an extreme auxotroph that lacks many central metabolic pathways (102,103). To get around this limitation, it parasitizes purines (104), amino acids (105), cholesterol (106, 107), long-chain fatty acids (108,109), carbon sources (110), and other metabolites (111) from the host. A. phagocytophilum is obligately intracellular and parasitizes amino acids and cholesterol from the host, in addition to manipulating host cell processes with secreted effectors (112)(113)(114)(115)(116)(117). From this perspective, both microbes cause stress to the host by competing for a finite amount of resources and disturbing normal cellular processes. Indeed, our evidence shows that tick-transmitted microbes stimulate the UPR and are restricted by its activity. Although cellular stress responses detect and respond to stress, they are not necessarily specific to types of stressors and instead respond by monitoring macromolecular threats to the cell (40,41,118). This more generalized signal widens the infection-sensing scope of possibility and reduces the requirement for an array of specific immune receptors. In this regard, a wide variety of stimuli would converge on a common immune outcome. Since the UPR is an evolutionarily conserved mechanism across eukaryotes (36)(37)(38), it is feasible that UPR-initiated immunity is a fundamental mode of pathogen sensing and host defense against a broad array of infectious organisms.
The absence of upstream IMD pathway regulators appears to be a shared trait among chelicerates and hemimetabolous insects (16-25, 27-33, 101, 120). Considering this observation, classically defined IMD pathway-initiating molecules may have evolved with holometabolous insects (those with complete metamorphosis), such as Drosophila and mosquitos, which undergo dramatic tissue remodeling between life stages (119, 121). The classically described IMD pathway is speculated to have arisen as a defense mechanism against microbiome-resident bacteria that are liberated with midgut breakdown during metamorphosis (31). In contrast, arthropods with incomplete metamorphosis, such as ticks and hemimetabolous insects, undergo some tissue remodeling during molting, but immature stages generally resemble adults in terms of morphology (119,122,123). Since these arthropods are phylogenetically more ancient than holometabolous insects (119,121,(124)(125)(126), it is possible that UPR activation of core IMD signaling molecules is ancestral to the classically defined pathway in Drosophila. Furthermore, our data support the idea that this network is conserved across arthropods. Specifically, IRE1a-mediated IMD pathway activation was observed both in Drosophila cells (Fig. S4B and C) and in two species of ticks ( Fig. 6 and 7). Collectively, this suggests that the UPR-IMD signaling network may have evolved in a common ancestor of hexapods and chelicerates.
In summary, we have discovered a linkage between cellular stress responses and arthropod immunity where the IRE1a-TRAF2 signaling axis initiates the IMD pathway (Fig. 8). The previous "orphaned" status of the IMD pathway in ticks was a perception that arose from comparative studies with the insect model organism Drosophila. This revelation underscores the importance of studying fundamental processes outside model organisms, which may be valuable for determining concepts that could be basally applicable across species. Our findings are conceptually important given that the IMD pathway widely impacts vector competence in many arthropods. With this commonality, one can envision a scenario where a conserved network across species may be an attractive target for future transmission intervention strategies.

MATERIALS AND METHODS
Bacteria and animal models. Escherichia coli cultures were grown in lysogeny broth (LB) supplemented with ampicillin at 100 mg mL 21 . Cultures were grown overnight at 37°C with shaking between 230 and 250 rpm.
Uninfected I. scapularis ticks were provided by the Biodefense and Emerging Infectious Diseases (BEI) Research Resources Repository from the National Institute of Allergy and Infectious Diseases (NIAID) (www.beiresources.org) at the National Institutes of Health (NIH) or from Oklahoma State University (Stillwater, OK, USA). Ticks were maintained in a 23°C incubator with 16/8-h light/dark photoperiods and 95 to 100% relative humidity. C3H/HeJ mice were purchased from The Jackson Laboratory, and C57BL/6 mice were obtained from colonies maintained at Washington State University. Six-to tenweek-old male mice were used for all experiments. C57BL/6 mice were infected intraperitoneally with 1 Â 10 7 host cell-free A. phagocytophilum organisms. C3H/HeJ mice were inoculated intradermally with 1 Â 10 5 low-passage-number B. burgdorferi. All mice were confirmed for infection status prior to tick placement by collecting 25 to 50 mL of blood from the lateral saphenous vein of each mouse 7 days postinfection. A. phagocytophilum burdens were enumerated by quantitative PCR (16s relative to mouse b-actin [128,129]). B. burgdorferi-infected blood was subcultured in BSK-II medium and examined for the presence of spirochetes by dark-field microscopy (130,131). Experiments involving mice were carried out according to guidelines and protocols approved by the American Association for Accreditation of Laboratory Animal Care (AAALAC) and by the Office of Campus Veterinarian at Washington State University (Animal Welfare Assurance A3485-01). The animals were housed and maintained in an AAALACaccredited facility at Washington State University in Pullman, WA. All procedures were approved by the Washington State University Biosafety and Animal Care and Use Committees.

FIG 8
The UPR triggers the IMD pathway in ticks. The tick-borne bacteria A. phagocytophilum and B. burgdorferi stimulate the UPR in I. scapularis ticks. IRE1a is activated by phosphorylation (P) and pairs with TRAF2. This signaling axis induces the IMD pathway, Relish activation, and antimicrobial responses that restrict pathogen colonization.
Plasmid construction. Both Ixodes IRE1a and TRAF2 were codon optimized for expression in human cell lines (GenScript). Primers (Table S1) were used to amplify full-length I. scapularis ire1a for cloning into pCMV/hygro-Negative Control Vector (SinoBiological; CV005) with HindIII sites. Full-length I. scapularis traf2 was amplified and cloned into pCMV-HA (New MCS) vector (received as a gift from Christopher A. Walsh; Addgene plasmid number 32530) using XhoI and EcoRV. All constructs were confirmed by sequencing (Eurofins Genomics).
Coimmunoprecipitation assay. Ixodes IRE1a-FLAG and TRAF2-HA expression was validated by immunoblotting whole-cell lysates with anti-FLAG-HRP (Sigma; A8592; 1:500) and anti-HA (Pierce; 26183; 1:1,000). After protein expression was confirmed, cross-linked agarose beads (anti-FLAG M2 [Sigma; A2220] and anti-HA [Pierce; 26181]) were washed twice with TBS (50 mM Tris, 150 mM NaCl; pH 7.5) and incubated with lysis buffer at 4°C for 1 h. Approximately 1 to 2 mg of cell lysate was combined with 80 mL (packed volume) of cross-linked agarose beads and incubated overnight at 4°C. Beads were washed 3 times with TBS, and protein was eluted by boiling in 50 mL of 4Â Laemmli buffer for 5 min. Protein interactions were evaluated by immunoblotting as described above.
Template-based homology modeling of Ixodes TRAF2 and the RNase/kinase domain of IRE1a. A BLAST search in the Protein Data Bank (PDB) using the Ixodes TRAF2 sequence returned the candidate template crystal structure of the TRAF-C domain from human TRAF2 (39.64% sequence identity). The human TRAF2 crystal structure (PDB code 1CA9) was used as a reference for building the homology model of the TRAF-C domain and part of the coiled-coil domain for Ixodes TRAF2 (residues 176 to 357) in SWISS-MODEL (71,134). QMEANDisCo was used to obtain a quality score, which defines how well the homology model aligns to reference structures in the PDB. Scores closer to 1 indicate that the homology model matches other reference structures well (135). Quality assessment of the TRAF2 homology model in QMEANDisCo gave a score of 0.69. The GalaxyRefine server was used to then further refine the Ixodes TRAF2 homology model, which increased the quality score in QMEANDisCo to 0.71 (136).
A PDB BLAST search for Ixodes IRE1a returned the candidate template crystal structure of the RNase/ kinase domain from human IRE1a (62.20% sequence identity). A homology model for the cytosolic RNase/kinase domain of tick IRE1a (residues 525 to 944) was built using the crystal structure of the RNase/kinase domain from humans (PDB code 6URC) with SWISS-MODEL (72,134). Quality assessment of the tick IRE1a homology model in QMEANDisCo gave a score of 0.78.
Prediction-driven docking of Ixodes TRAF2 and the RNase/kinase domain of IRE1a. A consensus interface predictor, CPORT (consensus prediction of interface residues in transient complexes), was used to assign residues at the interface of Ixodes TRAF2 and the IRE1a RNase/kinase domain (69). Predicted residues were used to define the docking interface between Ixodes TRAF2 and IRE1a for docking in HADDOCK 2.2 (74). The docked model was immersed in a solvent shell using the TIP3P water model, and a short 300-K molecular dynamics (MD) simulation was run to optimize side chains and improve interaction energetics (74). The cluster with the lowest Z score was chosen for further analysis. Docking models were then screened based on salt bridge interactions at the docking interface, and the model with the best chemical complementarity was used in the final analysis. PyMOL version 2.2.3 was used for all distance measurements of salt-bridge interactions (,4-Å cutoff) (PyMOL molecular graphics system; Schrodinger, LLC).
ROS assay. ISE6 cells were seeded at a density of 1.68 Â 10 5 cells per well in a black-walled, clear-bottom 96-well plate (Thermo Scientific; 165305) with L15C-300 medium. The cells were maintained under the growth conditions described above for the length of the experiments. All wells were treated for 1 h with 10 mM 29,79-dichlorofluorescein diacetate (DCF-DA; Sigma; D6883) in Ringer buffer (155 mM NaCl, 5  Pharmacological treatments, RNAi silencing, and qRT-PCR. ISE6 cells were seeded at 1 Â 10 6 cells per well and DAE100 cells were seeded at 5 Â 10 5 cells per well in a 24-well plate and pretreated with KIRA6, thapsigargin, or tunicamycin for indicated times and concentrations prior to infection. Cells were infected with A. phagocytophilum (ISE6) or A. marginale (DAE100) at an MOI of 50 for 18 h before collection in TRIzol (Invitrogen; 15596026). For DAE100 experiments, all incubations occurred at 34°C in a BD Campy Bag with no GasPak. RNA was extracted using the Direct-zol RNA microprep kit (Zymo; R2062). cDNA was synthesized from 300 to 500 ng total RNA with the Verso cDNA synthesis kit (Thermo Fisher Scientific; AB1453B). Bacterial burden and gene silencing were assessed by qRT-PCR with iTaq universal SYBR green Supermix (Bio-Rad; 1725125). Cycle conditions were as recommended by the manufacturer.
For transfection experiments, siRNAs and scrambled controls (scRNAs) were synthesized following directions from the Silencer siRNA construction kit (Invitrogen; AM1620). siRNA or scRNA (3 mg) was used to transfect 1 Â 10 6 ISE6 cells overnight with 2.5 mL of Lipofectamine 2000. Cells were infected with A. phagocytophilum (MOI, 50) for 18 h before being collected in TRIzol. RNA was isolated and transcripts were quantified by qRT-PCR as described above. All data are expressed as means and SEM.
Ex vivo I. scapularis and D. andersoni organ culture. Ten male and female unfed adult I. scapularis ticks were surface sterilized with continuous agitation in 10% benzalkonium chloride (Sigma; 12060) for 10 min, washed twice with sterile water, dried on sterile filter paper under aseptic conditions, and transferred to a sterile tube. Midgut and salivary glands were excised on a microscope slide in a pool of sterile 1Â PBS with 100 IU mL 21 penicillin and 100 mg mL 21 streptomycin (Gibco; 15140122). Tissues were placed in individual wells of a 96-well plate (Costar; 3595) with 100 mL of L15C-300 and incubated at 32°C with 1% CO 2 . Tissues were treated with 1 mM KIRA6 or 1% DMSO for 1 h before the addition of 1 Â 10 6 A. phagocytophilum. At 24 h postinfection, samples were collected following the addition of 100 mL of TRIzol. Tissues were homogenized using QIAshredder columns (Qiagen; 79654) according to the manufacturer's instructions prior to RNA extraction and qRT-PCR analysis, performed as described above.
Twenty male unfed adult D. andersoni ticks were surface sterilized and dissected as described above. Tissues were placed in individual wells of a 96-well plate with 100 mL of L15B. Tissues were pretreated with KIRA6 or vehicle control (DMSO) as stated above prior to the addition of 1 Â 10 6 A. marginale for 22 h. Samples were collected and processed as described above with qRT-PCR standard curves. All data are expressed as means and SEM.
RNAi silencing in nymphs and larvae. I. scapularis nymphs were microinjected as described previously (32,35). Ten-microliter Drummond microdispensers (DrummondSci; 3000203G/X) were drawn to fine-point needles using a Narishige PC-100 micropipette puller. I. scapularis nymphs were microinjected with 25 nl of siRNA or scRNA (;1,000 ng/mL) into the anal pore using a Drummond Nanoject III nanoliter injector (DrummondSci; 3000207). Ticks were allowed to rest overnight before being placed between the ears and on the back of an infected mouse. Each group was placed on a single mouse and fed to repletion (5 to 7 days). Nymphs were flash frozen with liquid nitrogen, individually crushed with a plastic pestle, and suspended in TRIzol for RNA extraction.
I. scapularis larvae were prechilled at 4°C for 5 min. Approximately 150 larvae were transferred to a 1.5-mL tube with 40 to 50 mL of either siRNA or scRNA (;1,000 ng/mL). The tubes were centrifuged at 3,000 Â g for 5 min to encourage submersion of the larvae in the dsRNA and were then incubated overnight at 15°C. The following day, ticks were dried and rested overnight before being placed on mice to feed until repletion (3 to 7 days). Larvae were flash frozen in liquid nitrogen and individually crushed with a plastic pestle. TRIzol was added before proceeding to RNA isolation and qRT-PCR analysis, performed as stated above.
xbp1 PCR and agarose gel electrophoresis. RNA was isolated from both ISE6 cells or replete I. scapularis nymphs (uninfected, A. phagocytophilum infected, or B. burgdorferi infected). ISE6 cells were treated with either 0.5 mM thapsigargin or A. phagocytophilum at an MOI of 50. Cells were collected 1, 3, 8, and 24 h posttreatment in TRIzol. RNA was isolated and cDNA synthesized as previously described. The cleavage status of xbp1 was assessed via PCR using DreamTaq Green PCR Mastermix (Thermo Scientific; K1082) and the xbp1 primers (Table  S1) with the cycling protocol recommended by the manufacturer. Samples were analyzed using a 3% agarose (Thermo Fisher; BP160) gel in 1Â Tris-borate-EDTA (TBE; Thermo Fisher; BP1333) with 0.5 mg mL 21 of ethidium bromide (Thermo Fisher; BP102) and imaged with a ProteinSimple AlphaImager HP system.
UPR and IMD gene expression profiling. Untreated I. scapularis nymphs were fed to repletion on A. phagocytophilum-infected mice or uninfected mice and frozen. The expression levels of UPR genes were assessed in individual ticks by qRT-PCR as previously described. Primers specific for bip, ire1a, xbp1, and traf2 were used (not shown). Data are expressed as means and SEM.
D. melanogaster S2* cells (1 Â 10 6 ) were seeded in Schneider's medium with 1 mM 20-hydroxyecdysone to prime the IMD pathway, as previous reported (137). Cells were treated with indicated concentrations of thapsigargin for 6 h or with 10 mM KIRA6 for 1 h prior to infection with A. phagocytophilum (MOI, 50) or B. burgdorferi (MOI, 50) for 6 h. Samples were collected in TRIzol, and RNA was isolated. IMD pathway-and Toll pathway-specific AMPs were quantified by qRT-PCR with primers as described above.
Statistical analysis. In vitro experiments were performed with 3 to 5 replicates. In vivo experiments involved the use of 10 to 20 ticks. Data were expressed as means and SEM and analyzed with either unpaired Student's t test or Welch's t test. Calculations and graphs were created with GraphPad Prism version 9.0. A P value of ,0.05 was considered statistically significant.

SUPPLEMENTAL MATERIAL
Supplemental material is available online only.

ACKNOWLEDGMENTS
We are grateful to Ulrike Munderloh (University of Minnesota) for providing ISE6 and DAE100 tick cell lines, to Jon Skare (Texas A&M Health Science Center) for providing B. burgdorferi B31 (MSK5), and to BEI Resources and Oklahoma State University for Ixodes scapularis ticks, and we are grateful for Addgene plasmid 32530, which was a gift from Christopher A. Walsh.