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A Streamlined Method for Transposon Mutagenesis of Rickettsia Parkeri

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A Streamlined Method for Transposon Mutagenesis of Rickettsia Parkeri bioRxiv preprint doi: https://doi.org/10.1101/277160; this version posted March 8, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission. 1 A streamlined method for transposon mutagenesis of 2 Rickettsia parkeri yields numerous mutations that 3 impact infection 4 5 Rebecca L. Lamason1,#a,*, Natasha M. Kafai1,#b, and Matthew D. Welch1* 6 7 8 1 Department of Molecular and Cell Biology, University of California, Berkeley, Berkeley, 9 CA 10 #a Current address: Department of Biology, Massachusetts Institute of Technology, 11 Cambridge, MA 12 #b Current address: Medical Scientist Training Program, Washington University in St. 13 Louis School of Medicine, St. Louis, MO 14 15 16 17 18 * Co-corresponding authors 19 E-mail: [email protected], [email protected] 20 1 bioRxiv preprint doi: https://doi.org/10.1101/277160; this version posted March 8, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission. 21 Abstract 22 The rickettsiae are obligate intracellular alphaproteobacteria that exhibit a complex 23 infectious life cycle in both arthropod and mammalian hosts. As obligate intracellular 24 bacteria, Rickettsia are highly adapted to living inside a variety of host cells, including 25 vascular endothelial cells during mammalian infection. Although it is assumed that the 26 rickettsiae produce numerous virulence factors that usurp or disrupt various host cell 27 pathways, they have been challenging to genetically manipulate to identify the key 28 bacterial factors that contribute to infection. Motivated to overcome this challenge, we 29 sought to expand the repertoire of available rickettsial loss-of-function mutants, using an 30 improved mariner-based transposon mutagenesis scheme. Here, we present the 31 isolation of over 100 transposon mutants in the spotted fever group species Rickettsia 32 parkeri. These mutants targeted genes implicated in a variety of pathways, including 33 bacterial replication and metabolism, hypothetical proteins, the type IV secretion 34 system, as well as factors with previously established roles in host cell interactions and 35 pathogenesis. Given the need to identify critical virulence factors, forward genetic 36 screens such as this will provide an excellent platform to more directly investigate 37 rickettsial biology and pathogenesis. 2 bioRxiv preprint doi: https://doi.org/10.1101/277160; this version posted March 8, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission. 38 Introduction 39 The Rickettsia are a genus of obligate intracellular alphaproteobacteria that are 40 divided into four groups - the spotted fever group (SFG), typhus group (TG), ancestral 41 group (AG), and transitional group (TRG) [1]. They inhabit arthropods (ticks, fleas, and 42 mites), and many can be transmitted to humans and other mammals. Pathogenic 43 species primarily target endothelial cells in the vasculature, causing a variety of vascular 44 diseases such as typhus and Rocky Mountain spotted fever [2]. Despite the prevalence 45 of rickettsial diseases throughout the world, we know little about the bacterial factors 46 required for infection and pathogenesis. 47 The SFG species Rickettsia parkeri, a tick-borne pathogen that causes a mild 48 form of spotted fever in humans [3,4], is emerging as a model organism to study SFG 49 rickettsial pathogenesis. R. parkeri can be studied under BL2 conditions and has animal 50 models of pathogenesis that mimic aspects of human infection [5,6]. Furthermore, the 51 R. parkeri life cycle closely matches that of the more virulent SFG species R. rickettsii 52 [7,8], the causative agent of Rocky Mountain spotted fever. Like its more virulent 53 relative, R. parkeri invades non-phagocytic cells and is taken into a primary phagocytic 54 vacuole [9]. They then break out of this vacuole and enter the cytosol to replicate and 55 grow [10]. R. parkeri and many other Rickettsia species also hijack the host cell actin 56 cytoskeleton to polymerize actin tails and undergo actin-based motility [11-13]. During 57 spread, motile R. parkeri move to a host cell-cell junction but then lose their actin tails 58 before entering into a short (~1 µm) plasma membrane protrusion that is subsequently 59 engulfed into the neighboring cell. The bacterium then lyses the double-membrane 60 secondary vacuole to enter the neighboring cell cytosol and begin the process again 3 bioRxiv preprint doi: https://doi.org/10.1101/277160; this version posted March 8, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission. 61 [14]. Because of its experimental tractability and the fact that its lifecycle is 62 indistinguishable from more virulent species, R. parkeri provides an attractive system for 63 investigating rickettsial host-pathogen interactions. 64 As an obligate intracellular pathogen, R. parkeri must produce virulence factors 65 that usurp or disrupt various host cell pathways. However, due to their obligate growth 66 requirement, it has been challenging to genetically manipulate the rickettsiae to identify 67 the key bacterial factors that contribute to infection [15]. Fortunately, recent advances 68 have expanded the genetic toolkit that can be used in the rickettsiae, allowing us to 69 more directly peer into the molecular mechanisms that drive rickettsial biology. Chief 70 among these advances was the development of a Himar1 mariner-based transposon 71 system for random mutagenesis of rickettsial genomes [16]. To date, smaller-scale 72 mutagenesis studies have been completed in the TG species R. prowazekii [16-18] and 73 the SFG species R. rickettsii [18,19]. 74 Despite these advances, we still do not know all of the critical bacterial factors 75 that mediate interactions with the host. Moreover, many of the genes in R. parkeri are 76 annotated to encode hypothetical proteins, which limits our ability to rationally explore 77 their functions. Therefore, we set out to expand the repertoire of available R. parkeri 78 mutants using a forward genetic screen. We used the mariner-based transposon 79 system [16] and developed a more streamlined protocol to rapidly isolate R. parkeri 80 mutants that alter plaque size. To date, we have isolated over 100 mutants that disrupt 81 genes predicted to function in a variety of pathways. We have previously published our 82 detailed analysis of three mutants – in sca2, rickA, and sca4 [14,20]. Here, we present 4 bioRxiv preprint doi: https://doi.org/10.1101/277160; this version posted March 8, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission. 83 the full panel of mutants to demonstrate the potential and ease of developing rickettsial 84 transposon libraries. 85 86 Materials and methods 87 Cell lines 88 Vero cells (monkey, kidney epithelial) were obtained from the University of 89 California, Berkeley tissue culture facility and grown in Dulbecco’s modified Eagle’s 90 medium (DMEM) (Invitrogen) containing 5% fetal bovine serum (FBS) at 37˚C in 5% 91 CO2. 92 93 Transposon mutagenesis in R. parkeri 94 R. parkeri Portsmouth strain was a gift from Dr. Chris Paddock (Centers for 95 Disease Control and Prevention). Wild-type R. parkeri were expanded and purified by 96 centrifugation through a 30% MD-76R solution, as previously described [14]. The 97 pMW1650 plasmid carrying the Himar1 mariner-based transposon [16] (a gift from Dr. 98 David Wood, University of South Alabama) was used to generate R. parkeri strains 99 carrying transposon insertions. To isolate small plaque mutants, a small-scale 100 electroporation protocol was designed. First, a T75 cm2 flask of confluent Vero cells was 101 infected with approximately 107 WT R. parkeri. Three days post-infection, when Vero 102 cells were at least 90% rounded, they were scraped from the flask. Infected cells were 103 spun down for 5 min at 1800 x g at 4˚C and resuspended in 3-6 ml K-36 buffer. To 104 mechanically disrupt infected cells and release bacteria, cells were either passed 5 bioRxiv preprint doi: https://doi.org/10.1101/277160; this version posted March 8, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission. 105 through a 27.5 gauge syringe needle 10 times, or vortexed at ~2900 rpm using a Vortex 106 Genie 2 (Scientific Industries Inc.) in a 15 ml conical tube containing 2 g of 1 mm glass 107 beads, with two 30 s pulses and 30 s incubations in ice after each pulse. This bead 108 disruption procedure was adopted for a majority of the screen because it was faster and 109 reduced the possibility of a needle stick. Host cell debris was pelleted for 5 min at 200 x 110 g at 4˚C. The supernatant containing R. parkeri was transferred to 1.5 ml 111 microcentrifuge tubes and spun for 2 min at 9000 x g at 4˚C. The bacterial pellets were 112 then washed three times in cold 250 mM sucrose, resuspended in 50 µl cold 250 mM 113 sucrose, mixed with 1 µg of pMW1650 plasmid, placed in a 0.1 cm cuvette, and 114 electroporated at 1.8 kV, 200 ohms, 25 µF, 5 ms using a Gene Pulser Xcell (Bio-Rad). 115 Bacteria were immediately recovered in 1.2 ml brain heart infusion (BHI) media. To 116 infect confluent Vero cells in 6-well plates, media was removed from each well, and 117 cells were washed with phosphate-buffered saline (PBS).
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