Adaptation of Anaplasma phagocytophilum to the tick vector is controlled by the transcriptional regulator Tr1

Adaptation of Anaplasma phagocytophilum to the tick vector is controlled by the transcriptional regulator Tr1 | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Adaptation of Anaplasma phagocytophilum to the tick vector is controlled by the transcriptional regulator Tr1 EricaRose Warwick , Rachel Burt , Jeffrey T. Badigian , Daniel Howell , Kyle T. Swallow , Chloe Leach , View ORCID Profile Azeza M. Falghoush , View ORCID Profile Dana K. Shaw , View ORCID Profile Ian T. Cadby , View ORCID Profile Jason M. Park doi: https://doi.org/10.1101/2025.11.12.688119 EricaRose Warwick 1 Department of Veterinary Microbiology and Pathology, Washington State University , Pullman, Washington, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Rachel Burt 2 Bristol Veterinary School, University of Bristol , Bristol, UK Find this author on Google Scholar Find this author on PubMed Search for this author on this site Jeffrey T. Badigian 1 Department of Veterinary Microbiology and Pathology, Washington State University , Pullman, Washington, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Daniel Howell 1 Department of Veterinary Microbiology and Pathology, Washington State University , Pullman, Washington, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Kyle T. Swallow 1 Department of Veterinary Microbiology and Pathology, Washington State University , Pullman, Washington, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Chloe Leach 1 Department of Veterinary Microbiology and Pathology, Washington State University , Pullman, Washington, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Azeza M. Falghoush 1 Department of Veterinary Microbiology and Pathology, Washington State University , Pullman, Washington, USA 3 College of Sciences, Sirte University , Libya 4 College of Medical Technology, Aljufra University , Libya Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Azeza M. Falghoush Dana K. Shaw 1 Department of Veterinary Microbiology and Pathology, Washington State University , Pullman, Washington, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Dana K. Shaw Ian T. Cadby 2 Bristol Veterinary School, University of Bristol , Bristol, UK Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Ian T. Cadby Jason M. Park 1 Department of Veterinary Microbiology and Pathology, Washington State University , Pullman, Washington, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Jason M. Park For correspondence: jpark12{at}wsu.edu Abstract Full Text Info/History Metrics Supplementary material Preview PDF ABSTRACT Rickettsial pathogens are strictly dependent on the cellular biology of their hosts for survival and replication. Predominantly transmitted by blood-feeding arthropods, these vector-borne pathogens are forced to adapt between the disparate environments of their mammalian host and arthropod vector. To achieve this, the Rickettsial bacteria Anaplasma phagocytophilum undergoes extensive transcriptional reprogramming with over 41% of its genes differentially transcribed between mammals and Ixodes scapularis ticks. How the bacterium orchestrates this dramatic transcriptional reprogramming is not understood. The gene tr1 encodes a Helix-Turn-Helix DNA-binding protein that is exclusively expressed during tick infection. Herein, we show that tr1 is essential for A. phagocytophilum survival in ticks and regulates the transcription of other genes necessary to adapt to the arthropod vector. We demonstrate that Tr1 is a DNA-binding protein and recognizes promotors of tick-specific genes in A. phagocytophilum, including secreted effector ateA , alternate components of type IV secretion system (T4SS), and membrane proteins. Our findings demonstrate that Tr1 is a master regulator of genes that are critical for A. phagocytophilum adaptation to the tick. INTRODUCTION Rickettsial pathogens are strictly dependent on the cellular biology of their hosts for survival and replication, which is reflected by their small genomes and reduced metabolic capacities. Instead these bacteria have evolved factors to mediate host-pathogen interactions and manipulate the intracellular environment 1 – 4 . These include specialized surface proteins mediating contact and uptake into cells 2 , 3 , 5 – 9 and effector molecules injected through secretion systems to redirect host cell pathways 10 – 23 . The large majority of rickettsial host-pathogen interactions are described in the context of mammalian infection 17 , 20 – 22 , 24 , 25 . However, since rickettsial bacteria are predominantly transmitted by blood-feeding arthropods, these findings only explore half of the pathogen lifecycle 26 . Mammals and arthropods are separated by over 680 million years of evolution and, as such, present distinct environments that Rickettsial pathogens must adapt to survive intracellularly 27 . The most common rickettsial pathogen in the United States, Anaplasma phagocytophilum , is transmitted to humans, domestic animals, and wildlife by the tick Ixodes scapularis 28 . While in the mammalian host, A. phagocytophilum infects and replicates within circulating neutrophils. In the tick, the bacteria initially infect the digestive tract and then migrate to the salivary glands, where they will survive through the molt 29 , 30 . In response to the disparate biology between the mammalian host and tick vector 31 , A. phagocytophilum undergoes extensive transcriptional reprogramming. Transcriptomic studies found that over 41% of A. phagocytophilum genes are differentially transcribed when comparing infected human monocytes and tick cells 32 , 33 . Indeed, Himar1 transposon disruption of host or vector specific A. phagocytophilum genes reduces bacterial survival in the respective human or tick cell cultures 10 , 34 – 37 . Together, this suggests transcriptional reprogramming of A. phagocytophilum is necessary for adaptation between the host and vector environments. While both transcriptomics and proteomics have shown A. phagocytophilum and other rickettsial pathogens undergo extensive retooling during tick infections 5 , 32 , 33 , 38 – 42 , the regulatory mechanisms that control these dramatic shifts are not known. One gene, tr1 , encodes a putative Helix-Turn-Helix DNA-binding protein (Tr1). tr1 was first noticed due to its proximity to outer membrane protein (OMP) genes omp_1X , omp_1N and the msp2 expression site 43 , 44 , but its impact on this neighboring operon is not defined. Strikingly, of all A. phagocytophilum genes tr1 has the highest transcriptional specificity for tick cell infection 32 , 33 . Herein, we demonstrate that tr1 is essential for A. phagocytophilum survival in tick cells, colonization of ticks in vivo , and expression of many genes necessary for adaptation to the tick. Further, we demonstrate that Tr1 is a DNA-binding protein and recognizes promotors of tick-specific genes in A. phagocytophilum, including secreted effector ateA , alternate components of type IV secretion system (T4SS), and membrane proteins. From this work we have identified Tr1 as a master regulator of genes that are critical for A. phagocytophilum adaptation to the tick. RESULTS Transcription of tr1 is specific to tick cell infection A. phagocytophilum HGE1 tr1 transcription was quantified and compared between bacteria cultured in the human monocyte-like HL60 cell line and embryonic tick ISE6 cells. Consistent with previous tiling array reports, tr1 transcription was >26 fold higher during growth in tick ISE6 cells over the human HL60 cell line ( Fig 1a ), validating that tr1 expression is highly specific to growth in tick cells 32 , 33 . Download figure Open in new tab Figure 1. tr1 transcription is specific to tick cell infection and necessary for A. phagocytophilum survival in tick cells. ( a ) tr1 transcription normalized to housekeeping gene rpoB during A. phagocytophilum culture in ISE6 tick cells and human HL60 cells. Bar indicate mean of four replicate infections each measured in two technical replicates show as points. ( b and c ) Growth of A. phagocytophilum tr1 ::Himar1 or control strain in cell culture infections of ( b ) human HL60 cells and ( c ) tick ISE6 cells. A. phagocytophilum burden measured by bacterial gDNA relative to eukaryotic host gDNA via qPCR. Data displayed as mean with ±SD of three biological replicates with two technical replicates each. Data are representative of three experimental replicates. * P < 0.05 (Mann-Whitney t -test). tr1 is necessary for A. phagocytophilum survival in tick cells The specificity of tr1 transcription to ISE6 cell culture prompted us to test if tr1 is necessary for A. phagocytophilum adaptation to ticks. To examine this possibility, we obtained a transposon mutant strain tr1 ::Himar1 from the previously published mutant collection 34 . An established control strain with the Himar1 transposon inserted in an intergenic non-coding region was used for comparison. This control strain is phenotypically comparable to wild-type in both mammalian and tick infection models 10 , 34 , 36 , 37 . tr1::Himar1 and the control A. phagocytophilum strain were purified from HL60 cell culture and used to infect either ISE6s or HL60s. Bacterial burdens were compared over time. During HL60 infection, the control and tr1 ::Himar1 strains grew equivalently ( Fig 1b ). However, during tick cell infection, tr1 ::Himar1 steadily declined ( Fig 1c ), indicating that it is required for A. phagocytophilum adaptation to tick cells. A. phagocytophilum requires tr1 to colonize ticks in vivo The in vitro tr1 ::Himar1 phenotype led us to ask if tr1 is similarly required in vivo for A. phagocytophilum adaptation to the tick. Mice were infected with either the intergenic control or the tr1 ::Himar1 mutant A. phagocytophilum . Seven days post infection bacterial burden in the mice was quantified and found to be equivalent between the strains, indicating tr1 is not required during murine infection ( Fig 2a ). Burden matched mice from these groups were used to feed I. scapularis larval ticks to repletion. We found ticks that fed on mice infected with tr1 ::Himar1 acquired significantly less A. phagocytophilum than those that fed on mice infected with the control strain ( Fig 2b ). These findings indicate that tr1 is dispensable for murine infection but required for colonization of the arthropod vector. Download figure Open in new tab Figure 2. tr1 is despicable during murine infection but essential for acquisition by ticks. ( a ) Mouse blood Anaplasma burden 7 days post intraperitoneal inoculation with 1 × 10 8 A. phagocytophilum tr1 ::Himar1 or control::Himar1 strains. Blood processed for gDNA and bacterial burden was measured by qPCR of A. phagocytophilum 16S rDNA versus mouse actin by ΔΔCt. Each strain was tested in twenty mice (½ male, ½ female). Each sample was tested in technical duplicate reaction. ( b ) Two burden-matched infected mouse pairs were used for Ixodes scapularis larvae infestation. Ticks were allowed to feed to repletion and detach. Whole replete I. scapularis larvae were processed for RNA. A. phagocytophilum bacterial loads were measured via qRT-PCR of A. phagocytophilum 16S rRNA levels versus mouse actin transcripts. Data includes ticks from two burden matched mouse pairs. From each mouse, 10–20 individual ticks were collected as biological replicates, and each qRT-PCR was performed in duplicate. * P < 0.005 (Welch’s t -test). Tr1 is a multimeric Helix-Turn-Helix DNA-binding protein Having established that Tr1 is important for adaptation to the tick host by A. phagocytophilum , we next used bioinformatics approaches to predict the functions of the Tr1 protein. First, a structural model of monomeric Tr1 protein (amino acids 1-183) was generated using ColabFold 45 . The resulting predicted Tr1 monomer model is comprised of two ordered and predominantly α-helical domains that are separated by a central disordered peptide linker and flanked by disordered peptide at both the N and C termini ( Fig 3a ). In total, the model contains approximately 58 residues of predicted disorder. Whilst the global confidence score for the predicted model was low (pTM=0.468), the local confidence scores for the two ordered domains were relatively high (pLDDT greater than 80 for residues ∼25-90 and ∼130-155, Supplementary figure 1). Download figure Open in new tab Figure 3. Structural modelling of the Tr1 protein. (a) Schematic of the predicted Tr1 protein domain architecture. Ordered domains are displayed as boxes and predicted disordered regions as solid black lines. (b) Cartoon representation of a predictive Tr1 dimer protein model. For one Tr1 chain the N-terminal DNA binding domain is colored plum, the C-terminal dimerization domain is colored light green, and disordered flanking and linker regions are colored pale yellow. The second Tr1 chain is colored grey. (c) Dimeric H-T-H domain of Tr1 (colored magenta) superimposed onto the crystal structure of HipB bound to its DNA operator from Shewanella oneidensis 76 (colored light blue; PDB: 4PU4). (d) Dimeric C-terminal domain of Tr1 (colored green) superimposed onto the crystal structure of EspR from M. tuberculosis 77 (colored brown; PDB: 4NDW) Next, we used the Dali server 46 to search the Protein Data Bank (PDB) for structures similar to Tr1, but which have been the subject of structure-function analyses. Amongst the results of the Dali search, nine of the ten structures most similar to Tr1 were DNA-binding proteins with similarity to the N-terminal domain of Tr1 and which function as dimers. Comparing the N-terminal domain of Tr1 with these structures reveal that Tr1 is likely to be a five-helix bundle which contains a DNA-binding helix-turn-helix motif (H-T-H), similar to cI/Cro-like transcription factors ( Fig 3a ). Prompted by this observation, we next used ColabFold Multimer to generate a model of dimeric Tr1 protein. In the resulting model, the N-terminal domains of Tr1 are closely associated with one another, forming a symmetrical dimer ( Fig 3b ). The C-terminal domains of the two Tr1 monomers, which each resemble two antiparallel helices separated by a turn, interdigitate with one another forming an additional point for dimerization between the two protein chains ( Fig 3b ) . Whilst the global confidence score for the dimeric model of Tr1 was higher than that of the monomeric model (dimer pTM 0.557), the confidence score for the interface of the dimeric Tr1 model was low (ipTM=0.545). We considered that the large proportion of disordered protein in Tr1 could skew the interface confidence scores in our predictions, so we tried to predict dimeric models for Tr1 residues corresponding to the N- and C-terminal domains only. Interface confidence scores for dimeric Tr1 N- or C-terminal domains in isolation were high enough to suggest plausible interactions (piTM=0.763 and 0.69, respectively). Structures of dimeric H-T-H proteins identified by our Dali search were superimposed onto the Tr1 dimer model to further assess this predicted structure. The N-terminal domain of Tr1 dimerizes in manner typical of other dimeric H-T-H proteins, including those bound to DNA ( Fig 3c ). Frequently, H-T-H proteins that bind DNA as symmetrical homodimers bind to palindromic DNA sequences with each protein binding to a half-site of the palindrome. From these structural comparisons, we predict this is likely to also be the case for Tr1. Fewer structural homologues of the C-terminal domain of Tr1 were identified but we noted similarity between the Tr1 dimer model and the EspR transcription factor of Mycobacterium tuberculosis ( Fig 3d ) . EspR dimerizes via a domain similar to the Tr1 C-terminal domain and this is required for EspR to bind DNA with high-affinity 47 . Next, we used gel filtration chromatography to assess whether purified recombinant Tr1 protein could form dimers or other multimers in solution. Recombinant Tr1 protein was purified and affinity tags removed prior to running on a gel filtration column. We reasoned that removing any additional protein sequences from Tr1 would yield protein most similar to that found in its native environment. Since the expected molecular weight of monomeric recombinant Tr1 is 21.3 kDa, we also ran purified MBP and GFP, predominantly monomeric proteins with respective molecular weights of 43.8 kDa and 28.5 kDa, as controls. Both MBP and GFP eluted from the gel filtration column as monodisperse peaks whereas Tr1 eluted as two overlapping peaks, indicating that Tr1 exists in multiple forms in solution ( Fig 4a ). SDS-PAGE analysis of fractions collected from these experiments confirmed that the peaks contained the expected proteins ( Fig 4b ). Apparent molecular weights for the peaks were calculated with a standard curve based on their elution volumes. The calculated apparent molecular weights for GFP and MBP were 31.6 and 45.7 kDa, respectively, roughly consistent with their expected weights as monomers. The second peak of Tr1 eluted at an apparent molecular weight of 79.4 kDa, suggesting that Tr1 can form tetramers in solution. The first peak of Tr1 eluted at a volume outside of our standard curve and potentially contained aggregated proteins. We generated a predicted tetrameric model of full-length Tr1, but global confidence scores were noticeably lower than those for other Tr1 models (pTM 0.414), so do not present this data here. Further biochemical data are required to guide additional predictive modelling of Tr1 multimers. Download figure Open in new tab Figure 4. Tr1 forms multimers in solution. (a) Chromatograms of Tr1, MBP, and GFP separated by gel filtration. The Tr1 protein lacks tryptophan residues and absorbs weakly at 280 nm so the peaks are shown as normalized absorbance. (b) SDS-PAGE gel of fractions collected from equivalent volumes in gel filtration experiments. The left-hand lane of each gel contains molecular weight markers with the known weights labelled. Taken together, we predict that Tr1 is comprised of H-T-H and dimerization/multimerization domains joined and flanked by regions of disorder. Comparisons indicate that Tr1 might have similarity to cI/Cro transcription factors and EspR from M. tuberculosis. Gel filtration experiments demonstrate that Tr1 forms multimers in solution, supporting these predictions, however, a high confidence predicted model of a Tr1 tetramer could not be generated Tr1 binds the promotors of neighboring genes omp1X and omp1N The tr1 gene was first noticed in studies examining expression of downstream genes omp1X , omp1N , and the msp2/p44 expression site 43 , 44 ( Fig 5a ). We tested Tr1 binding upstream of tr1 , omp1X , omp1N , and the msp2/p44 expression locus by electrophoretic mobility shift assays (EMSA) using DNA probes for sequences preceding each gene ( Fig 5a ). EMSA shifts indicated Tr1 complexed with promotors p- tr1 ( Fig 5b ), p- omp1X ( Fig 5c ) and p- omp1N ( Fig 5d ). At higher Tr1 concentrations p- tr1 and p- omp1N also displayed secondary complexes suggesting multiple Tr1 binding sites or higher order Tr1 oligomer complexes ( Fig 5b,d ). DNA sequence upstream of the msp2 expression locus did not shift at any concentration of the Tr1 protein, indicating Tr1 does not individually regulate msp2 ( Fig 5e ). Download figure Open in new tab Figure 5. Tr1 binds promotors of neighboring omp1X and omp1N. ( a ) Diagram of tr1 gene with downstream neighboring outer membrane protein genes omp1X , omp1N , and the msp2/p44 expression site. ( b through e ) EMSA shifts with increasing rTr1 (0, 0.0625, 0.125, 0.25, 0.5, and 1μM ) with DNA probes of ( b ) tr1 , ( c ) omp1X , ( d ) omp1N , and ( e ) msp2/p44 promotor sequences. ( f and g ) Transcription of omp1X and omp1R from control or tr1 ::Himar1 A. phagocytophilum mutant strains at 24 hours post infection in ( f ) human HL60 or ( g ) tick ISE6 cells. Transcripts measured by qRT-PCR and normalized to rpoB via ΔΔCt. Data displayed as mean with ±SD of four replicate infections measured with two technical replicates each. * P < 0.05 (Mann-Whitney t -test). ( h ) msp2/p44 transcripts totals across all gene variants sequenced by RNAseq from tr1 ::Himar1 or control A. phagocytophilum infected ISE6 tick cells. Bars are mean ±SD of four independent infections, each shown as points. n.s. > 0.05 (Mann-Whitney t -test). To ask how loss of tr1 affected these genes’ transcription, RNA was collected from control and tr1 ::Himar1 A. phagocytophilum infected ISE6 cells. Surprisingly, qPCR measuring omp1X and omp1N transcription from tr1 ::Himar1 and control A. phagocytophilum strains ( Fig 5f,g ) found that only omp1N displayed a small but significant expression difference during tick cell infection ( Fig 5g ), suggesting Tr1 binding alone does fully account for the behavior of these genes. tr1 is required for A. phagocytophilum transcriptional shift during tick cell infection Tr1’s predicted role as a transcriptional regulator 43 , 48 and the inability of the tr1 ::Himar1 to survive in tick cells led us to ask how the loss of tr1 affects the transcriptome during tick cell infection using RNA sequencing (RNAseq). ISE6 tick cells were infected with A. phagocytophilum tr1::Himar1 or the control strain. Our previous qPCR for A. phagocytophilum gDNA from infected ISE6 cells found that the tr1 ::Himar1 strain persists up to day 3 before declining relative to the control strain ( Fig 1c ). Since measuring genomic DNA (gDNA) may also reflect dead bacteria, we measured A. phagocytophilum 16s RNA relative to I. scapularis actin transcripts as an indicator of bacterial survival at 12, 24, and 48 hours post infection (hpi). At 24 hours tr1 ::Himar1 survival was 63% of the control strain but fell to only 10% by 48 hpi ( Fig S2 ). To capture transcriptional differences when tr1 ::Himar1 bacteria remained viable we performed RNAseq 24 hpi. Genes downstream of tr1 , omp1X , omp1N , and msp2 43 ( Fig 5a ) were all measured in the RNAseq data. Similar to the qRT-PCR measurements ( Fig 5g ), omp1X did not significantly differ between tr1 ::Himar1 and the control strain. omp1N had a small but significant increase in expression relative to the control ( Table S1 ). Measuring msp2 transcription is complicated by the presence of multiple msp2 pseudogenes, which A. phagocytophilum uses to evade the mammalian antibody response by antigenic variation. For individual msp2 variants to be transcribed, the pseudogenes recombine into the expression site adjacent to omp1N 43 , 44 , 49 – 52 ( Fig 5a ). In our RNAseq data, 97 msp2 variants were detected, with 15 having significant transcription differences between tr1 ::Himar1 and the control strain ( Table S2 ). To quantify total msp2 transcription from the expression site, we summed all msp2 transcripts from the RNAseq data. This found no difference in overall msp2 transcription between the tr1 ::Himar1 and control strain ( Fig 5h ). Expression differences among individual msp2 variants likely reflects msp2 diversity bottle neck during Himar1 library construction and/or msp2 recombination since the tr1 ::Himar1 and control strains were purified from the larger collection. These finding again suggest the importance of tr1 extends beyond its immediate genetic neighborhood. Beyond tr1 ’s immediate neighbors, RNAseq identified 177 genes as differentially expressed between tr1 ::Himar1 and the control A. phagocytophilum strain (p-adj2 fold upregulated in the tr1 ::Himar1, although none have known host or vector-specific expression 32 , 33 and are largely ribosomal or other core bacterial genes 34 . Conversely, 14 genes had >2 fold reduced transcription from the tr1 ::Himar1 mutant ( Table 1 ). Seven of the of the these, (HGE1_03907/APH_0916, ateA , tr1 , HGE1_01872/APH_0406, msp4 , HGE1_04767/APH_1111, HGE1_03162/APH_0720) have known tick-specific expression patterns 10 , 32 , 33 , which indicates that tr1 is needed to upregulate A. phagocytophilum genes specific for the arthropod vector. View this table: View inline View popup Download powerpoint Table 1. Genes with ≥ 2 fold reduced expression in tr1 ::Himar1 mutant during tick cell infection. Tr1 binds promotors of genes of tick-specific A. phagocytophilum genes We next asked if Tr1 binds promotors of any tick-specific genes impacted by the tr1 ::Himar1 mutation. EMSA were performed with increasing Tr1 protein against promotor sequences preceding; ateA , HGE1_06057/APH_1380, HGE1_03162/APH_0720, HGE1_03907/APH_0916, HGE1_01872/APH_0406, and msp4 ( Fig 6 ). Tr1 shifted probes for ateA ( Fig 6a ), HGE1_06057 ( Fig 6b ), and msp4 ( Fig 6c ) at all concentrations, with the majority of the probe band shifted at the highest. HGE1_03162 ( Fig 6d ), HGE1_03907/APH_0916 ( Fig 6e ), and HGE1_01872/APH_0406 ( Fig 6f ) probes also shifted at Tr1 concentrations ≥ 0.125 µM. Similar to EMSAs with probes for tr1 , omp1X and omp1N, the higher Tr1 concentrations produced secondary shift bands, suggesting higher order complexes. These findings demonstrate direct Tr1 interaction with promoters for tick-specific A. phagocytophilum genes. These EMSA results and differential expression of these genes by the tr1 ::Himar1 mutant strain indicates direct regulation by Tr1. Download figure Open in new tab Figure 6. Tr1 binds the promotors of tick-specific A. phagocytophilum genes. EMSA shifts with increasing rTr1 (0, 0.0625, 0.125, 0.25, 0.5, and 1μM) tested against promotor sequences of ( a ) ateA , ( b ) HGE1_06057 , ( c ) HGE1_01872 , ( d ) HGE1_03162 , ( e ) HGE1_03907 , and ( f ) mps4 . Tr1 regulated genes are important for A. phagocytophilum survival in tick cells Inability of the tr1 ::Himar1 to survive in tick cells and the reduced expression of known tick-specific A. phagocytophilum genes led us to ask if these genes are important for tick cell infection. We previously showed ateA ::Himar1 was similarly defective for survival in tick cells and acquisition by ticks 10 . From the A. phagocytophilum Himar1 mutant library, we identified and isolated three additional mutants disrupted in genes putatively regulated by Tr1: HGE1_01872 , HGE1_03162 , and HGE1_06057 . Growth of all mutant strains in human HL60 cell culture was comparable to the control strain ( Fig 7a ). Accordingly, we infected ISE6 tick cells with A. phagocytophilum strains tr1 ::Himar1, ateA ::Himar1, HGE1_01872 ::Himar1, HGE1_03162 ::Himar1, HGE1_06057 ::Himar1, and intergenic control::Himar1. At 8 days post infection all test strains had significantly reduced Anaplasma burden in the tick cells relative to the intergenic Himar1 control, with tr1::Himar1 being the most severe ( Fig 7b ). From this, we demonstrated three additional tick-specific genes HGE1_01872, HGE1_03162, and HGE1_06057 as critical for infecting ticks. Our results suggest that the tick-specific defect of tr1 ::Himar1 mutant strain stems from its failure to express genes controlled by Tr1 and critical for adaptation to the tick cell environment. Download figure Open in new tab Figure 7. Tr1 impacted genes are necessary for survival during tick cell infection. Survival of indicated Himar1 transposon mutant A. phagocytophilum strain relative to intergenic control::Himar1 strain in ( a ) human HL60 (48 hpi) and tick ISE6 cells (8 dpi). Bacterial burden measured by qRT-PCR of A. phagocytophilum 16s vs Ixodes actin transcripts. Data displayed as mean ±SD of four replicate infections measured in two technical replicates each. Graph representative of three replicate experiments. * < 0.005 (Mann-Whitney t -test). tr1 is necessary for tick-specific alterations in the T4SS apparatus The promotor sequences of ateA and HGE1_06057 had high affinity for Tr1 by EMSA ( Fig 6 a,b ). Previously we identified AteA as the first A. phagocytophilum T4SS secreted effector specific to tick infection 10 . HGE1_06057 is also predicted as a T4SS substrate by multiple effector prediction algorithms 12 , 53 . Tr1 binding to promotors of effectors prompted us to ask if Tr1 influences expression of T4SS machinery itself. An oddity of the Anaplasma T4SS is that some components of the apparatus are encoded by multiple paralogs 16 , 23 . In particular, the needle like pilus of the A. phagocytophilum T4SS is encoded for by eight paralogs of the pilin gene virB2 ( Fig 8a ). Transcriptomics studies noticed different combinations of virB2 paralogs are upregulated when A. phagocytophilum infects mammalian or tick cells 32 , 33 . Because tiling arrays used in these studies can conflate signal among paralogs with close sequence identity 33 , we first validated virB2 expression. We designed primers specific to each virB2 paralog. We grew A. phagocytophilum in human HL60s or tick ISE6 cells and measured transcription of each virB2 paralog by qRT-PCR. As in the transcriptomic studies, mammalian cultured A. phagocytophilum only expressed virB2 paralogs virB2_1 and virB2_2 ( Fig 8b ). Conversely, transcription of virB2_1 and virB2_2 was reduced during growth in tick cells, while virB2 _3 → virB2_8 were upregulated ( Fig 8b ). To examine if the tick-specific expression pattern among the virB2 paralogs is dependent on tr1 we reviewed our RNAseq findings for the tr1 ::Himar mutant. Both virB2_1 and virB2_2 expression were significantly elevated (1.47 and 1.38 fold) in the tr1 ::Himar1. While not significant due to low number of total reads virB2_6 and virB2_7 had reduced expression (0.55 and 0.218 fold) in the tr1 ::Himar1 relative to the control strain ( Table S1 ). Together this indicated that the tr1 ::Himar1 strain fails to adjust expression among the virB2 paralogs during tick cell infection and retains a virB2 expression pattern similar to mammalian infection. Download figure Open in new tab Figure 8. Tr1 is required for tick-specific remodeling of the T4SS pilus. ( a ) Diagram of the virB2 paralog genes (locus HGE1_04947 – HGE1_04867), the location of the virB2_5 ::Himar1 insertion mutation (red), and potential promotor regions subject to EMSA (green). ( b ) Transcription of the virB2 genes during wild-type A. phagocytophilum infection of mammalian HL60 and tick ISE6 cells. Data shown as mean ±SD of four biological replicates. ( c,d,e ) EMSA shifts with increasing rTr1 (0, 0.0625, 0.125, 0.25, 0.5, and 1μM ) with putative promotor sequences upstream of ( c ) virB2_1 , ( d ) virB2_6 , and ( e ) virB2_8 . ( f and g ) Growth of A. phagocytophilum virB2_5 ::Himar1 or control strain in cell culture infections of ( f ) human HL60 cells and ( g ) tick ISE6 cells. A. phagocytophilum burden measured by bacterial gDNA relative to eukaryotic host gDNA via qPCR. Data displayed as mean with ±SD of three biological replicates with two technical replicates each. Data are representative of three experimental replicates. * P < 0.05 (Mann-Whitney t -test). Tr1 binds promotors of T4SS virB2 paralogs differentially expressed in tick infection We asked if Tr1 can directly bind promotor regions among the differentially expressed virB2 paralogs. DNA probes designed upstream of virB2_1 , virB2_6, and virB2_8 ( Fig 8a ) were tested with increasing concentrations of Tr1 protein by EMSA. Tr1 shifted the promotors of both virB2_1/2 ( Fig 8c ) and virB2_6/7 ( Fig 8d ), with virB2_1/2 shifting completely at the highest concentration. As with other targets, possible secondary complexes were also visible, suggesting possible higher order interactions. Minimal shifting was seen with probes designed upstream of virB2_8 ( Fig 8e ) suggesting either independent regulation, or co-regulation with the upstream genes virB_6/7 . Our findings indicate that Tr1 directly participates in regulation of tick-specific expression patterns of T4SS components. Tick-specific virB2 paralogs are necessary for A. phagocytophilum survival in tick cells As with the other Tr1 regulated targets, we asked if the tick-specific virB2 paralogs are necessary for A. phagocytophilum survival in tick cells. From the transposon mutant library, we obtained a mutant strain disrupted in virB2_5 ( virB2_5 ::Himar1) ( Fig 8a ). In the A. phagocytophilum HGE1 strain genome (APHH00000000) virB2_5 is not annotated and therefore was not initially mapped by RNAseq. However, we do detect transcription from the virB2_5 locus during tick cell infection ( Fig 8b ). The virB2_5 ::Himar1 mutant grew as well as the control strain in human HL60 cells ( Fig 8f ). A. phagocytophilum virB2_5 ::Himar1 and the intergenic control::Himar1 strain were collected from HL60 cell culture and used to infect ISE6 tick cells. Eight days post infection Anaplasma burden of virB2_5 ::Himar1 was greatly reduced relative to the control::Himar1 strain in the tick cells ( Fig 8e ). This finding indicates that tick-specific virB2 paralogs are important for A. phagocytophilum adaptation to the tick. Tr1 homologs are found throughout the Rickettsiales Having established a role for Tr1 in the survival of A. phagocytophilum within the tick host, we queried whether Tr1 homologs are present in other Rickettsiales 43 , since many of these bacteria also have life cycles that involve cycling between vertebrate and arthropod hosts. Using sequence-based searches within the Rickettsiales, we identified candidate homologs of Tr1 in diverse Anaplasmataceae genomes. The tr1 genes of Anaplasma , Ehrlichia , and Neoehrlichia share a high degree of genomic synteny and are invariably co-localized with ndk (encoding nucleoside-diphosphate kinase) and one or more OMP-encoding genes. In Wolbachia and “Candidatus Mesenet “ the tr1 locus is more variable with ndk being lost and OMP-encoding genes being co-localized at the tr1 locus in some, but not all, species. We used Foldseek to extend our search for more distant Tr1 homologs, filtering our results for Rickettsiales bacteria. Using this approach, we identified Tr1 proteins in Rickettsia and “Candidatus Tisiphia “ species. Whilst the tr1 locus in Rickettsia species is not syntenic with those from the Anaplasmataceae, the Rickettsia tr1 gene is most often co-localized with sca4. Intriguingly, sca4 has been identified as having a role in fitness of Rickettsia parkeri within the tick host 54 . Yet more distant Tr1 homologs were also identified in Midichloria and Orientia , although in the latter the C-terminal domain is truncated or lost. Candidate tr1 homologues are co-localized with genes that encode proteins with unknown functions in many Rickettsiales. These results indicate that Tr1 represents a family of transcription factors found broadly across the Rickettsiales, suggesting that some of our findings could be extrapolated to other bacteria of this Order. DISCUSSION As a vector-borne pathogen, survival in both the tick and the mammal is essential to complete the transmission cycle of A. phagocytophilum and be maintained in the environment. We know the bacteria undergoes extensive transcriptional changes between the two environments 32 , 33 and have identified individual adaptations that are critical for either mammalian or tick infection 34 – 37 . Understanding how rickettsial pathogens switch their host tropism to the arthropod, could inform interventions to disrupt the transmission cycle. In this work we identified Tr1 as a critical switch regulating A. phagocytophilum adaptation to the tick and identified Tr1-controlled genes necessary for survival in the arthropod. Collectively our findings uncover a central node in a network of changes necessary for rickettsial bacteria to complete their vector-borne lifecycle. Due to the obligate intracellular lifecycle and lack of compatible extrachromosomal plasmids, genetic manipulation of A. phagocytophilum remains challenging. The Himar1 transposon library has proven to be an invaluable resource, allowing the first phenotypic testing of gene disruptions in Anaplasma 34 . The library was generated in HL60 cell culture, which prohibits mutation in genes essential for survival in mammalian cells. However, A. phagocytophilum genes specific to tick infection were readily mutated 34 , 35 . Such mutants have been leveraged to uncover tick-specific roles of the T4SS effector AteA 10 , duplicate T4SS component VirB6-4 36 , and surface protein modifying O-methyltransferase 37 . In our study we extensively relied on mutants from this library to uncover tick-specific contributions of tr1 and five of the genes it impacts. Until more targeted genetic tools are available in this system, the A. phagocytophilum Himar1 transposon library remains a critical resource. Initial interest in the tr1 gene stemmed from its location upstream of outer membrane proteins genes omp1X , omp1N , and msp2/p44 expression site 43 , 44 . ApxR was identified as the first Anaplasma DNA-binding transcriptional regulator, and was later shown to bind its own promotor and upstream region of the p44/msp2 expression site. Similarly, the ApxR homolog EcxR in the related rickettsial pathogen Ehrlichia chaffeensis , binds promotors of tr1 and downstream outer membrane protein genes 55 , 56 . E. chaffeensis Tr1 also binds upstream membrane protein genes p28 and omp-1B 55 . We found A. phagocytophilum Tr1 binds DNA sequences upstream of omp1X and omp1N . However, only expression of omp1N differed between the tr1 ::Himar1 mutant and control strain, indicating tr1 is not the only factor controlling these genes. Though the number, identity, and arrangement of adjacent membrane protein genes downstream of tr1 in Anaplasma and Ehrlichia differ greatly 44 , 55 , common models may emerge showing how Tr1 and ApxR/EcxR participate in regulation of the tr1 adjacent surface protein genes. Looking past its immediate chromosomal neighbors multiple tick-specific membrane protein genes had reduced expression in the Himar1:: tr1 and were directly bound by Tr1. These include; HGE1_03162 ( Aph_0720 ), HGE1_01872 ( Aph_0406 ), msp4 , and HGE1_03907 ( Aph_0916 ). Transcription of all four is highly specific to A. phagocytophilum infection in tick cells 32 , 33 . Reflecting the expression pattern, we found HGE1_03162 and HGE1_01872 are necessary for tick cell infection. Adjacent to HGE1_01872, A. phagocytophilum encodes two HGE1_01872 paralogs asp62 ( HGE1_01862, Aph_0404 ) and asp55 ( HGE1_01867, Aph_0505 ). Antibodies against Asp62 and Asp55 were found to limit A. phagocytophilum infection of human HL60 cells 57 . While Asp62 and Asp55 are expressed during both mammalian and tick cell infection, HGE1_01872 is specific to the arthropod vector 32 , 33 . Both HGE1_01872 and Msp4 are substrates of an O-methyltransferase required for tick infection and promote binding to Ixodes cells 37 , 58 , 59 . Similar to HGE1_01872, HGE1_03907 (Aph_0916) is adjacent to the invasion protein gene aipA , which is necessary for infection of mammalian cells 60 , suggesting HGE1_03907 represents a arthropod-specific AipA alternative. These findings implicate Tr1 as the switch regulator governing this remodeling of the A. phagocytophilum ’s surface proteome necessary for tick infection. Beyond surface protein genes, Tr1 is playing a broader role controlling genes expressed during infection of the tick. Our findings identify Tr1 as a critical regulator of tick-specific effector AteA and specialization of the T4SS VirB2 pilus necessary for adaptation to the tick. The full repertoire of effectors secreted by A. phagocytophilum is unknown, but effector predicting algorithms propose as many as 49 12 , 53 . So far, few have been experimentally examined 17 , 21 , 22 , 24 , 61 – 63 , with only AteA investigated in context of the arthropod vector 10 . The E. chaffeensis ApxR homolog, EcxR, was also shown to regulate components of the T4SS apparatus during mammalian cell infection 64 . However, EcxR has yet to be examined during tick infection, and binding to effector to virB2 pilin genes is untested. Uncovering the full regulome of Tr1 and how it works with other regulators to manage adaptation between the mammalian and tick environments will likely require larger unbiased binding and regulation studies. How does Tr1 recognize its DNA targets and regulate gene expression? Bioinformatic analysis indicates that the Tr1 protein has similarity to H-T-H transcription factors and, inferred from this and our protein modelling, is likely to dimerize and bind palindromic DNA motifs to exert influence on transcription on downstream genes. Consistent with this, we also identified a putative dimerization domain in the C-terminus of Tr1 and we observed that recombinant Tr1 protein forms multimers in solution, probably tetramers and possibly other higher order structures. Other H-T-H proteins such as the λ repressor, share this ability to form multimers of dimers, enabling them to bind to multiple DNA sites co-operatively. Indeed, multiple shifts of Tr1-DNA complexes in our EMSAs, supporting that binding of Tr1 might be co-operative at some of its target sites. Understanding if or how the multimerization of Tr1 is regulated during infection, the interplay between Tr1 and other transcription factors at promoter regions, and the identification of a consensus DNA target site and regulon will be crucial for determining how Tr1 mediates A. phagocytophilum adaptation to the arthropod host. We identified candidate Tr1 homologs throughout the Rickettsiales. In both Anaplasma and some Rickettsia, the Tr1 gene is co-localized with genes that are either differentially expressed between the tick and vertebrate host or have been implicated in survival in tick cells. This functional conservation of the tr1 loci between these two species makes it interesting to consider that Tr1 might have a role in transcriptional remodeling that underpins the cycling between vertebrate and arthropod hosts in diverse Rickettsiales. However, this leaves the question of what the role of Tr1 in Rickettsiales that survive exclusively within arthropods, such as Wolbachia, is. Furthermore, in many Rickettsiales, the tr1 gene is co-localized with genes of no known function. Whilst genomic synteny suggests that Tr1 fulfills similar roles in Anaplasma, Ehrlichia and Neoehrlichia, further study is required to determine how Tr1 functions across diverse Rickettsiales. Altogether, our work identifies Tr1 as an essential master regulator that remodels the A. phagocytophilum transcriptome to infect the tick vector. Further, we identified multiple Tr1 regulated genes that contribute to tick cell infection. With most studies focusing on how A. phagocytophilum infects mammals, this work provides insights into an understudied aspect of pathogen biology, uncovering wide-reaching transcriptional events that underpin adaption to the arthropod vector. Unable to be transmitted vertically to host progeny, A. phagocytophilum must cycle between the host and tick vector to be maintained in the environment. Tr1 and the genes under its control are an essential part of this system. Uncovering how A. phagocytophilum and other rickettsial bacteria achieve this is key to developing interventions to interrupt the transmission cycle. MATERIALS AND METHODS Bacterial and eukaryotic cell culture Escherichia coli used for plasmid construction, plasmid amplification, and protein expression was cultured with solid and liquid lysogeny broth (LB) medium with the addition of kanamycin or zeocin 25 µg ml −1 antibiotics as needed to select for plasmid encoded resistance genes. For production of recombinant Tr1 proteins, E. coli strain DE3 (New England Biolabs) was used with LB or Terrific Broth (TB) supplemented with 100 µg ml −1 kanamycin. HL60 human promyelocytic cells (ATCC; CCL-240) were maintained in Roswell Park Memorial Institute (RPMI) 1640 medium with 10% FBS and 1× Glutamax. HL60 cultures were kept at 37°C with 5% CO 2 in a humidified incubator. HL60 cell cultures were limited to < 20 passages to prevent phenotypic drift and kept between 5 × 10 4 and 1 × 10 6 cell/ml to prevent differentiation. Wild-type A. phagocytophilum strain HGE1 and HGE1 derived Himar1 transposon mutants were grown in HL60 cells as previously described 34 . All Himar1 mutant strains were obtained from the previously established A. phagocytophilum mutant collection 34 . Status of A. phagocytophilum infections in HL60 cells was monitored by Diff-Quick Romanowsky–Giemsa staining. To generate host-cell-free A. phagocytophilum, peak infected HL60 cultures (>95% infected cells) were gently sonicated to disrupt HL60 cell membranes and liberated bacteria were separated from host cell debris by differential centrifugation. Anaplasma numbers were estimated as previously described 65 , 66 . I. scapularis (Say) embryonic tick cells (ISE6) were maintained in L15C-300 medium with 10% FBS (Sigma; F0926), 10% tryptone phosphate broth (TPB; BD; B260300) and 0.1% lipoprotein cholesterol concentrate (MP Biomedicals; 219147680) 67 , in sealed tissue culture treated flasks, and incubated at 34°C 68 . Infected ISE6 cell cultures were additionally supplemented with 0.25% NaHCO 3 and 25 mM HEPES buffer (Sigma), cultured in vented flasks, and kept at 34°C in a humidified chamber with 4% CO 2 . A. phagocytophilum mutant growth curves A. phagocytophilum burden in HL60 and ISE6 cells was evaluated as previously described 36 , 37 . Briefly, HL60 cells were seeded to 24 well plates at 5×10 4 cells/well and infected at an MOI of 1. Three to four replicate wells were harvested at the time of inoculation and tested time post inoculation. ISE6 cells were seeded to 24 well plates at 3×10 5 cells/well. After ISE6 cells adhered to the plate for at least 6 hours, they were inoculated at an MOI of 50 with host-cell-free A. phagocytophilum purified from HL60 cultures. Bacteria were allowed to infect the tick cells for 4 hours, after which the media was exchanged three times to remove excess extracellular bacteria. Four replicate wells were collected post media exchanged and subsequent time points. Pelleted samples were frozen and later processed for RNA or gDNA using Zymogen Quick-RNA Micro-prep or QIAGEN DNeasy blood and tissue kits. Ratios of bacterial and host cell gDNA copies was measured by qPCR as previously described 10 , 36 , 37 Levels of live A. phagocytophilum Himar1 mutant bacteria in each infected ISE6 or HL60 cell sample was measured by qRT-PCR targeting A. phagocytophilum 16S rRNA and I. scapularis or human β-actin transcripts ( Table S3 ) and compared relative to the intergenic Himar1 control strain by ΔΔCt 10 , 65 , 69 . Measuring A. phagocytophilum gene transcription HL60 and ISE6 cells in 24 well plates were infected with wild-type A. phagocytophilum HGE1 as described above. At 24 hours post infections four wells were collected and processed for RNA Zymo Quick-RNA micro-prep kit ® (ZymoResearch) according to product protocols for tissue culture samples. cDNA was generated with Verso cDNA Synthesis Kit (ThermoFisher). Transcripts of A. phagocytophilum genes of interest were measured by qPCR using gene specific primers ( Table S3 ) and SYBR green iTaq universal Supermix (Bio-Rad; 1725125) according to Bio-Rad specified cycle conditions. Genes of interest were measured relative to housekeeping gene rpoB and compared between human and tick cell infections by ΔΔCt. Animal infection All mice were purchased as six weeks of age from The Jackson Laboratory. Gender balanced groups of ten C57BL/6 mice were infected as previously described 10 with either the tr1 ::Himar1 or the intergenic Himar1 mutant A. phagocytophilum . The intergenic Himar1 mutant was previously shown to be phenotypically equivalent to wild-type 10 , 36 , 37 . Blood was collected seven days post inoculation and A. phagocytophilum burden in the blood was measured by qPCR as in prior studies (16S rRNA / mouse β-actin 10 , 65 , 69 ) (Table S3). Larval I. scapularis ticks purchased from Oklahoma State University (Stillwater, OK, USA) were kept at 23°C with >95% humidity and a 16/8-h light/dark photoperiods. Mice were selected from the intergenic Himar1 and tr1 ::Himar1 strain infected mice with equivalent Anaplasma burden. Each mouse was infested with 200 naïve larval I. scapularis and allowed to feed to repletion. Detached replete ticks were individually processed and assayed for A. phagocytophilum burden by qRT-PCR measuring 16S rRNA versus I. scapularis β-actin transcripts (Table S3), compared by absolute quantification 10 , 65 , 69 . All animal use protocols were approved by the Washington State University Institutional Animal Care and Use Committee (ASAF #6630) and animal housing facilities at Washington State University in Pullman, WA maintains AAALAC-accreditation. RNA sequencing ISE6 cells were seeded to 12 well plates at 5×10 5 cells/well and allowed to adhere for 6 hours. The intergenic transposon control and Himar1:: tr1 mutant A. phagocytophilum strains were prepared as host-cell-free bacteria from >95% infected HL60 cultures immediately before ISE6 infection. Adhered ISE6 cells were infected at an MOI of 100. Twenty-four hours post infection media was exchanged three times to remove excess extracellular bacteria, and four replicate wells were collected and RNA isolated with Zymogen Quick-RNA Mini-prep kits. RNA was submitted to the WSU Genomics Core for total RNA-seq analysis. Sequencing libraries were prepared using Tru-seq Stranded Total RNA kit, with Ribo-Zero Plus and libraries, sequenced using a HiSeq 2500, and results exported as FASTQ files as previously discribed 70 . RNA-seq data were aligned with the A. phagocytophilum HGE1 genome (GCF_000478425.1) 71 . Transcript quantification and differential gene expression were analyzed by HTSeq and DESeq2 72 . Overexpression and purification of recombinant Tr1 protein The tr1 open reading frame, codon optimized for expression in E. coli, was synthesized and cloned by Twist Bioscience into a modified pET28a expression vector. The resultant plasmid encoded the Tr1 protein fused at its N-terminus to a hexa-histidine tagged maltose binding protein (MBP) with a tobacco etch virus (TEV) protease recognition site between Tr1 and MBP. The plasmid was transformed into E. coli DE3 for overexpression of the recombinant Tr1 protein. Briefly, overnight cultures, grown at 37 °C in selective LB media, were used to inoculate flasks of TB. Over-expression of MBP-Tr1 was induced by the addition of 0.5 mM IPTG once cultures had reached an OD 600 of 0.5-0.6 and then grown for an additional 18 hours at 25 °C. Cells were harvested by centrifugation and kept at -20 °C for short-term storage. To purify MBP-Tr1 protein, cell pellets were resuspended in Buffer A (20 mM imidazole, pH 8.0; 20 mM Tris-HCl, pH 8.0; 400 mM NaCl; 2 mM beta-mercaptoethanol) and then lysed by sonication. Lysates were clarified by centrifugation and then loaded onto a HisTrap HP purification column (Cytiva). The column was washed extensively with Buffer A prior to elution of the His-tagged MBP-Tr1 with stepwise (10, 20, 100%) washes with Buffer B (as Buffer A but with 400 mM imidazole). Fractions containing MBP-Tr1 were pooled, concentrated by centrifugal-filtration, then further purified by gel-filtration on a Superdex 200 column (Cytiva) equilibrated in SEC Buffer (20 mM Tris-HCl pH 8.0; 250 mM NaCl; 2 mM beta-mercaptoethanol). Fractions containing MBP-Tr1, which eluted as a single peak from the gel-filtration column, were supplemented with his-tagged TEV protease and incubated at 4 °C for 16 hours to cleave the His-tagged MBP protein from Tr1. The resulting protein preparation was passed over a HisTrap HP column which was then washed step (5, 10, 100%) with Buffer B. Tr1 protein typically passed through the HisTrap HP column or eluted in 5% Buffer B washes and were thus separated from the His-tagged TEV and MBP proteins, which eluted in 100% Buffer B. This purification strategy yielded highly pure Tr1 protein lacking any tags. Gel filtration Purified tagless Tr1 protein, green fluorescent protein (GFP) and maltose binding protein proteins were analyzed by gel filtration using a HiPrep Sephadex 75 16/600 column (Cytiva), using Buffer B. GFP and MBP were expressed using pET vectors (modified pET41c and pET28a, respectively) in E. coli DE3 and purified by nickel affinity chromatography or amylose affinity chromatography using standard methods. Electrophoretic-Mobility-Shift-Assays Promoter proximal DNA sequences for use in Electrophoretic-Mobility-Shift-Assays (EMSAs) were amplified by PCR. Primers for this purpose were designed to yield DNA fragments comprising the ATG start codon of a selected gene plus 250 bp of upstream DNA sequence. In each case, the forwards primer was appended with an M13 sequence upstream of the primer sequence (5’- GAG CGG ATA ACA ATT TCA CAC AGG). The resulting DNA fragments were separated by agarose gel electrophoresis and fragments of the anticipated size were excised and purified (QIAquick Gel Extraction Kit). These purified fragments were then used as the templates for a secondary PCR using an M13 sequence primer labelled with FAM at the 5’ end in conjunction with the fragment-specific reverse primer used in the primary PCR. The resulting FAM-labelled PCR products were resolved by agarose gel electrophoresis and purified as above. Sequences of primers used are included in table S3 . EMSA assays were performed using a modified protocol described elsewhere 73 . Briefly, for each DNA fragment an EMSA mixture comprising (100 ng FAM-labelled DNA fragment;200 ng of herring sperm DNA; 5% glycerol; 10 mM Trs-HCl, pH 8.0; 25 mM KCl) was prepared and 8 µl of this dispensed into a series of tubes. These were then supplemented with 2 µl of purified Tr1 protein at varying concentrations, or with buffer alone, and then mixed by gentle pipetting. Tr1 protein was used at final concentrations of 0.25, 0.5, 1, 2, and 4 µM for all EMSAs presented here. Resulting mixtures were incubated at 25 °C for 30 minutes before being loaded onto 7.5 % polyacrylamide gels buffered with 0.25x TBE and 5% glycerol. Gels were run for 60 minutes at 100 V and then fluorescent images (green channel) were collected on a ChemiDoc MP Imaging System (Bio-Rad). Bioinformatics Protein structures were analyzed and visualized in Coot 74 and ChimeraX 75 . Modelling and the generation of predicted structures were done using ColabFold and AlphaFold Server with searches for homologues in the PDB being achieved with the Dali Server 45 , 46 . Download figure Open in new tab Figure S1. Predicted Local Distance Difference Test (pLDDT) and Multiple Sequence Alignment (MSA) plots from ColabFold of Tr1. (a) pLDDT plot shows that local confidence in the Tr1 model is high in the N- and C-terminal domains, supporting the proposal that Tr1 is comprised of two ordered domains. (b) MSA plot showing that the Tr1 N- and C-terminal domains are evolutionarily conserved. Download figure Open in new tab Figure S2. Early survival of tr1 ::Himar1 strain during tick cell infection. A. phagocytophilum tr1::Himar1 or control strain in cell culture infections of tick ISE6 cells. A. phagocytophilum burden measured by bacterial 16s relative to tick actin transcripts via qRT-PCR. Data displayed as mean with ±SD of three biological replicates. Data are representative of three experimental replicates. *P < 0.05 (Mann-Whitney t-test). FUNDING This work was funded through generous support from; the National Institutes of Health (NIAID grant numbers R21AI178392, R21AI154023, R21AI151412, R61AI179933, R01AI162819), Washington State University Intramural College of Veterinary Medicine grants program funded by the Marvel Shields Autzen Fund and Stanley L. Adler Research Fund, and Washington State University, College of Veterinary Medicine. The Cadby laboratory was also funded by Royal Society Research Grant RGS\R1\231117 and Academy for Medical Sciences Springboard Award SBF009\1204. AUTHOR CONTRIBUTIONS EricaRose Warwick: Methodology, Investigation, Visualization, Reviewing & Editing, R achel Burt: Methodology, Investigation, Visualization, Jeffrey T. Badigian: Methodology, Investigation, Daniel Howell: Methodology, Investigation, Kyle T. Swallow: Methodology, Investigation, Chloe Leach: Methodology, Azeza M. Falghoush: Methodology, Dana K. Shaw: Methodology, Investigation, Review & Editing. Ian T. Cadby: Conceptualization, Supervision, Methodology, Investigation, Writing, Visualization, Funding acquisition. Jason M. Park: Conceptualization, Supervision, Methodology, Investigation, Writing, Visualization, Funding acquisition. ACKNOWLEDGEMENTS We are grateful to Ulrike Munderloh, Lisa Price and Nicole Burkhardt at the University of Minnesota (Saint Paul, MN) and Kelly Brayton at Washington State University (Pullman, WA) for providing strains from the Himar1 transposon and methodological guidance. Kelly Brayton for sharing resources. Deirdre Fahy provided administrative support. Washington State University (WSU) Genomics Core performed RNA-sequencing and analyses. Funder Information Declared National Institutes of Health, https://ror.org/01cwqze88 , R21AI178392 , R21AI154023 , R21AI151412 , R61AI179933 , R01AI162819 Marvel Shields Autzen Fund , na Stanley L. 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