Impact of Active Metabolism on Chlamydia trachomatisElementary Body Transcript Profile and Infectivity

Scott Grieshaber,a Nicole Grieshaber,a Hong Yang,b Briana Baxter,a Ted Hackstadt,c Anders Omslandb

aDepartment of Biological Sciences, College of Science, University of Idaho, Moscow, Idaho, USAbPaul G. Allen School for Global Animal Health, College of Veterinary Medicine, Washington State University,Pullman, Washington, USA

cLaboratory of Bacteriology, Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases,Hamilton, Montana, USA

ABSTRACT Bacteria of the genus Chlamydia include the significant human pathogensChlamydia trachomatis and C. pneumoniae. All chlamydiae are obligate intracellular para-sites that depend on infection of a host cell and transition through a biphasic develop-mental cycle. Following host cell invasion by the infectious elementary body (EB), thepathogen transitions to the replicative but noninfectious reticulate body (RB). Differentia-tion of the RB back to the EB is essential to generate infectious progeny. While the EBform has historically been regarded as metabolically inert, maintenance of infectivityduring incubation with specific nutrients has revealed active maintenance of the infec-tious phenotype. Using transcriptome sequencing, we show that the transcriptome ofextracellular EBs incubated under metabolically stimulating conditions does not clusterwith germinating EBs but rather with the transcriptome of EBs isolated directly from in-fected cells. In addition, the transcriptional profile of the extracellular metabolizing EBsmore closely resembled that of EB production than germination. Maintenance of infec-tivity of extracellular EBs was achieved by metabolizing chemically diverse compounds,including glucose 6-phosphate, ATP, and amino acids, all of which can be found in ex-tracellular environments, including mucosal secretions. We further show that the EB celltype actively maintains infectivity in the inclusion after terminal differentiation. Overall,these findings contribute to the emerging understanding that the EB cell form is activelymaintained through metabolic processes after terminal differentiation to facilitate pro-longed infectivity within the inclusion and under host cell free conditions, for example,following deposition at mucosal surfaces.

IMPORTANCE Chlamydiae are obligate intracellular Gram-negative bacteria that are re-sponsible for a wide range of diseases in both animal and human hosts. According tothe Centers for Disease Control and Prevention, C. trachomatis is the most frequently re-ported sexually transmitted infection in the United States, costing the American healthcare system nearly $2.4 billion annually. Every year, there are over 4 million new cases ofChlamydia infections in the United States and an estimated 100 million cases worldwide.To cause disease, Chlamydia must successfully complete its complex biphasic develop-mental cycle, alternating between an infectious cell form (EB) specialized for initiatingentry into target cells and a replicative form (RB) specialized for creating and maintain-ing the intracellular replication niche. The EB cell form has historically been consideredmetabolically quiescent, a passive entity simply waiting for contact with a host cell toinitiate the next round of infection. Recent studies and data presented here demonstratethat the EB maintains its infectious phenotype by actively metabolizing a variety of nu-trients. Therefore, the EB appears to have an active role in chlamydial biology, possiblywithin multiple environments, such as mucosal surfaces, fomites, and inside the host cellafter formation.

Received 5 February 2018 Accepted 1 May2018

Accepted manuscript posted online 7 May2018

Citation Grieshaber S, Grieshaber N, Yang H,Baxter B, Hackstadt T, Omsland A. 2018. Impactof active metabolism on Chlamydiatrachomatis elementary body transcript profileand infectivity. J Bacteriol 200:e00065-18.https://doi.org/10.1128/JB.00065-18.

Editor Yves V. Brun, Indiana UniversityBloomington

Copyright © 2018 American Society forMicrobiology. All Rights Reserved.

Address correspondence to Scott Grieshaber,scottg@uidaho.edu, or Anders Omsland,anders.omsland@wsu.edu.

RESEARCH ARTICLE

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KEYWORDS Chlamydia, Chlamydia trachomatis, gene expression, intracellularbacteria, intracellular parasites, metabolism

The genus Chlamydia includes bacterial obligate intracellular parasites of both humansand animals. Chlamydia trachomatis, the most clinically significant human-adapted

species, is comprised of over 15 distinct serovars which cause sexually acquired infections,as well as the ocular infection trachoma, the leading cause of preventable blindness indeveloping countries (1). All chlamydiae undergo a characteristic biphasic developmentalcycle that includes physiologically specialized cell forms. Upon host cell entry by theinfectious elementary body (EB), morphological transition to the reticulate body (RB) isessential for propagation. Completion of the developmental cycle by differentiation back tothe EB form is required for generation of infectious progeny.

The EB and RB cell forms exhibit clear and contrasting biological and physiologicalspecialization. The EB is nonreplicative, has highly condensed chromatin, and is theonly cell form capable of initiating entry into host cells upon contact. In part based onexperimental data supporting the “energy parasite” hypothesis presented by Molder in1962 (2), the classical view of the EB is that of a metabolically “inert” entity (3). This viewhas largely been retained despite demonstration that EB infectivity depends on activetype III secretion (4–6). In contrast to the EB, the RB is replicative, larger with relaxedchromatin, and incapable of establishing infection (7). The chlamydial developmentalcycle is initiated when the EB cell type mediates entry into the host cell. After entry, theEB undergoes dramatic changes in gene expression profile coincident with morpho-logical differentiation to the RB (8). This early transitional gene expression leads to theestablishment of a unique replicative niche, the chlamydial inclusion. Upon establish-ment of the inclusion, the RBs begin to replicate (9). To ensure regeneration of EBs, theRBs must change gene expression to an EB-promoting pattern. This process is asynchro-nous; some RBs differentiate to the EB cell type as early as 18 h postinfection (serovar L2),while others continue replicating until they also eventually differentiate (9, 10).

The biology of the EB cell type is incompletely understood. That host cell-freemetabolic activity by the EB form of the environmental organism Protochlamydiaamoebophila and C. trachomatis can be detected by Raman microspectroscopy (11) andthat EB metabolic activity of the C. trachomatis EB can be stimulated by specificmetabolites (12, 13) suggest the EB cell type across the phylum Chlamydiae may playa larger role in active regulation of chlamydial pathogenesis than is generally appre-ciated. Similarly, infectivity of the C. trachomatis EB also declines more rapidly in theabsence of nutrients (12). Metabolism may also play a role in the EB-to-RB transition. EBgermination requires release of the compacted nucleoid structure by dissociation of thehistone like proteins HctA and HctB. This disassociation is in part regulated throughmetabolic activity as an intermediate in the isoprenoid biosynthetic pathway triggershistone release from the chromosome (14).

This study investigated the biological role of EB metabolism after terminal RB-EBdifferentiation. Based on previous work demonstrating that the EB can utilize glucose-6-phosphate (G6P) to synthesis protein and ATP, and with the EB simply definedphenotypically as the infectious form of C. trachomatis, we tested the central hypoth-esis that metabolic activity in the EB affects/dictates EB phenotype. The EB canmetabolize a variety of nutrients, and EB metabolism is necessary for optimal mainte-nance of infectivity. Our data demonstrate that, regardless of the nutrient conditiontested, the transcriptional profile of the metabolizing EBs more closely resembles theprofile of EB formation than EB germination. The induction of EB metabolism prolongedthe infectious phenotype regardless of the metabolites available, suggesting that the C.trachomatis EB phenotype can be maintained by access to chemically diverse metab-olites, including amino acids, G6P, and ATP. Surprisingly, although we could detecttranscription and protein synthesis in EBs, protein synthesis was not required formaintenance of infectivity. The ability of the EB to actively maintain the infectiousphenotype was also evident inside the inclusion. EBs in the inclusion mimicked the

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behavior of EBs under axenic conditions in that they were capable of protein synthesisbut did not require protein synthesis to remain infectious for up to 24 h after formation.

The ability of EBs to actively maintain their infectivity over an extended time periodvia chemically diverse substrates lends additional support to the existence of undera-ppreciated biological complexity in the chlamydial infectious cycle.

RESULTSGerminating intracellular EBs and EBs metabolizing extracellularly exhibit

distinct transcription profiles. The EB is triggered to undergo robust protein and ATPsynthesis in the absence of a host cell when incubated in the nutrient medium CIP-1(13). CIP-1 consists of intracellular phosphate buffer (IPB), nutrients (fetal bovine serum[FBS], amino acids, nucleotides [GTP, CTP, UTP, and ATP], and G6P), and the reducingagent dithiothreitol (DTT). Our previous studies additionally demonstrated that EBspreferentially used G6P over ATP as an energy source for de novo protein synthesis (13).To further understand the role of nutrients and metabolism in EB biology, the effectsof metabolism on gene expression patterns was assessed by transcriptome sequencing(RNA-seq). To this end, the transcriptional response of density gradient (DG) purifiedEBs to incubation under axenic conditions was compared to the response duringinfection of HeLa cells. The quality of the EB stock used for RNA-seq was characterizedby electron microscopy (EM), the ability to respond as previously determined to G6P,and by determining the ratio of genome equivalents (GE) to infectivity, the lattermeasured by an inclusion-forming unit (IFU) assay. The stock EBs incorporated[35S]cysteine-methionine in a dose-dependent manner (see Fig. S1 in the supplementalmaterial) (13), and EM micrographs showed that the inoculum consisted primarily ofuniform EB-sized cells with condensed nucleoids (see Fig. S2 in the supplementalmaterial). The GE/IFU ratio was calculated to be 89% � 2% standard errors of the mean(SEM), which is comparable to previously published data for C. trachomatis L2 (13, 15).To compare the transcriptomic responses, stock EBs were incubated in CIP-1 (0.5 mMG6P) or modified versions containing different concentrations of G6P (0 or 0.05 mM) for6 h, or used to infect cultured HeLa cells for 2, 5, 12, 18, and 24 h. Infection time pointsrepresent important biological transitions in the chlamydial developmental cycle (Fig.1). In the case of C. trachomatis serovar L2, germination and differentiation of the EB tothe RB occurs between 0 and 6 h postinfection, while RB replication is initiated around8 to 11 h and continues for the remainder of the infection (9). Differentiation of the RBto the EB occurs asynchronously starting at about 18 h postinfection and continuesuntil the end of the cycle (9, 10) (Fig. 1). To avoid changes in gene expression duringchlamydial purification, infected HeLa cells were processed for total RNA directly andchlamydial specific RNA enriched as described in Materials and Methods. For the earlytime points (2, 5, and 12 h), cells were incubated with a high multiplicity of infection(MOI; �1,000) and washed with heparin containing Hanks balanced salt solution (HBSS)

FIG 1 Schematic of the timing of the C. trachomatis L2, Bu434 developmental cycle. The developmental cycle ofC. trachomatis spans approximately 48 h and can be defined according to the timing of developmental transitionsbetween EB and RB cell forms. Germination, here defined as the time from EB internalization to the presence withinthe inclusion of fully mature and replication-competent RBs, occurs within 6 h postinfection (hpi). RB replication ensuesin the absence of EB generation until approximately 18 hpi, from which time point asynchronous RB-EB differentiationcan be observed, characterized by the presence of both RBs and EBs within the same inclusion (9, 10).

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after 5 min. Heparin washes reverse the electrostatic attachments of EBs that have notcommitted to cell entry (16). Early inclusions appear as separate vacuoles and developindependently until fusion occurs at about 12 h postinfection (17). After entry thegerminating EBs modify these early inclusions, recruit the dynein motor protein, andtraffic along microtubules to the microtubule organizing center (MTOC) of the cell (16).Since this is a phenotypic readout for germination, we measured the trafficking of ourinoculum by staining parallel infected coverslips and assaying for inclusion traffickingto the perinuclear region of the infected cells since this process is dependent on EBgermination, transcription, and translation (16, 18). Trafficking was quantified by mea-suring the percentage of perinuclear fluorescent signal compared to the fluorescencein entire cells. At 5 h postinfection, 93% � 1.5% of the signal was perinuclear,suggesting a uniform population of early inclusions (see Fig. S3 in the supplementalmaterial). HeLa cells were infected at an MOI of �5 for the 18- and 24-h samples. Thetranscriptome of the purified EB stock described above and a purified RB sample werealso included in the RNA-seq analysis. The purified RB sample was isolated from HeLacells at 17 h postinfection by density gradient (DG) centrifugation and evaluated by EM.The EM micrographs show that the RBs were uniform in size, were larger than the EBs,and did not contain visible EBs (see Fig. S2 in the supplemental material).

Biological replicates of each treatment were subjected to RNA-seq using the Illuminaplatform to determine the global transcriptome. Transcript levels were normalizedusing DESeq2 which were then analyzed by principal-component analysis (PCA) to plotsample-to-sample variance (19). This analysis showed that the transcription profile ofEBs incubated extracellularly, regardless of G6P concentration, clustered more closelywith the transcription profile of the EB inoculum than that of EBs germinating in hostcells (Fig. 2A). DESeq2 was used to calculate the log2 fold change in transcript levels foreach gene for each treatment compared to the EB inoculum (see Tables S1 to S19 in thesupplemental material). The 10 most highly induced genes in the 2-h HeLa and CIP-1samples were plotted in a heatmap ordered from high to low values in reference to theHeLa 2-h sample (Fig. 2B). The CIP-1 condition (0.5 mM G6P) was chosen as arepresentative of all G6P treatments as all showed a similar transcription profile. Nineof the top ten genes induced during EB germination in HeLa cells were lower in EBsincubated in CIP-1 (Fig. 2B), with the exception of CTL0699, encoding a predictedinclusion membrane protein (Inc) of unknown function (20). The reciprocal was alsotrue in that all 10 of the highest induced genes in CIP-1 were lower in intracellularlygerminating EBs (Fig. 2B). Hierarchical clustering of the DESeq2 transcription data wasalso used to compare expression patterns of genes related to specific cellular processes.The similarities of expression patterns for transcripts involved in cell division, DNAreplication, and protein synthesis were visualized using Python’s Seaborn clusteringvisualization tools (21) (Fig. 2C, D, and E, respectively; genes listed in Table S20 in thesupplemental material). These gene groups were derived from a combination ofontology generated through automated annotation and ontology generated by ourannotation based on published data. The genes included in the ontology groupingswere based on predicted functional pathways and not on expression profiles. In everycase the overall transcriptional profile of EBs incubated in CIP-1 clustered away from EBsgerminating in a host cell (Fig. 2C to E). Of note is that the 24-h time point clusteredcloser to the 5-h time point for cell division and genes involved in DNA replication.Because the inclusion contains both EBs and RBs at the 24-h time point, the heterog-enous population likely leads to the mixed clustering of transcript profiles noted at thistime point.

To assess changes in the expression of transcripts involved in different biologicalroles during chlamydial development, we next focused on genes involved in either EBgermination or EB formation and identified genes that appeared to be uniquelyregulated under axenic conditions. EB development is reliably initiated at �18 hpostinfection for C. trachomatis serovar L2; therefore, EB formation genes were definedas those expressed at higher levels at 24 h postinfection compared to the 18-h timepoint (listed in Fig. S4A in the supplemental material). For a first comparison, log2 fold

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change was calculated for each gene in HeLa 24 h and the CIP-1 (0.5 mM G6P)treatments compared to the purified 18-h RB control. The changes in transcriptexpression of these genes were visualized using hierarchical clustering of the log2 foldchange values. The transcription levels of the majority of the EB formation genes werehigher at both 24 h (HeLa 24 h) and in the extracellular EBs (CIP-1, 0.5 mM G6P)compared to the RB control (see Fig. S4A in the supplemental material). The change intranscription of EB formation genes in response to CIP-1 compared to the EB inoculumwas also evaluated. Hierarchical clustering showed that EBs incubated in CIP-1 dis-played a mixed pattern of gene transcription, with most genes transcribed higher orwith little change compared to the EB inoculum. A few genes were transcribed at lowerlevels than the EB control (see Fig. S4A in the supplemental material). However, onlyCTL0306, CTL0307, CTL0071, and CTL0398 (all encoding hypothetical proteins of un-

FIG 2 The transcription profiles of extracellular metabolically activated EBs do not resemble EB germination. The effect of metabolic activity on EB physiologicalstate was assessed by analysis of transcription profile signatures. Purified EBs were incubated under axenic conditions in the presence of differentconcentrations of G6P, a preferred energy source for EBs, and the EB transcript profiles compared to that of bacteria at different stages during intracellulardevelopment. (A) PCA analysis visualizing differences in gene expression patterns between sample replicates and treatments. The PCA analysis showed thatthe transcript profile of activated EBs was closer to that of nonincubated EB controls than to germinating EBs. Apparent separation between the processes ofmetabolic activation and germination suggests the EB is actively maintained. (B) Expression profile of the top 10 induced genes in axenically incubated EBscompared to the top 10 induced genes in EBs isolated from HeLa cells at 2 h postinfection illustrates clearly distinct transcriptional programs are activated underthe respective conditions. (C to E) Comparisons of the expression patterns between sample treatments of genes related to specific physiological processes wereevaluated using cluster analysis. The transcriptional pattern of genes related to cell division (C), DNA replication (D), and protein synthesis (E) show that theexpression pattern of EBs undergoing extracellular metabolic activation cluster away from bacteria, including EBs, at different stages during infection of HeLacells. Taken together, these results show that EBs incubated under axenic conditions in the presence of G6P maintain the transcriptional signatures ofinfection-competent EBs rather than initiating transcriptional programs consistent with germination.

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known function) and glgA were expressed between 3- and 6-fold lower compared tothe EB control (P � 0.01 for all genes).

Germination-related genes were defined as those with transcript levels that did notchange or were expressed higher at 5 h than 2 h in HeLa cells and were expressedhigher than the EB control (i.e., induced at both 2 and 5 h). The use of the 5-h samplecompared to the 2-h sample was to reduce the effect of carryover transcripts sincethese decrease upon infection and continue to decrease between 2 and 5 h postin-fection (8). Cluster analysis indicated that the majority of germination-related genes inCIP-1-incubated samples had transcript levels significantly lower or at similar levels tothe EB control; only CTL0673 was upregulated by at least 2-fold (2.1-fold, P � 0.01; seeFig. S4B in the supplemental material).

To identify genes that were uniquely expressed (noncanonical regulation) at thetranscriptional level under extracellular conditions, we again used hierarchical cluster-ing. Noncanonically regulated genes were defined as those with transcript levelsinduced in CIP-1 compared to the EB control, but lower in HeLa cells at 24 h comparedHeLa cells at 18 h (i.e., likely not involved in EB production) (see Fig. S3C in thesupplemental material). Three major types of genes were notable in this group, namely,type II secretion genes, amino acid transport and synthesis genes, and a subset of Incgenes (see Fig. S4 in the supplemental material).

Taken together, these data suggest that the overall transcription profile of extra-cellular metabolizing EBs more closely resembles that of fully developed EBs ratherthan EBs germinating to the RB form. However, there is a group of genes whosetranscription profiles appear not to follow this pattern. The functional grouping of thesegenes does not readily suggest a single dysregulated pathway and may instead reflectthe transcriptional profile of EBs responding to a unique environment.

Active maintenance of EB infectivity in an extracellular environment. Collec-

tively, our data suggest that the EB is capable of responding to metabolic signals toproduce a transcriptional response distinct from that representing EB germination. TheEB cell type is specialized for host cell invasion in order to initiate the next round ofreplication and maintain the infectious cycle. Therefore, we tested whether EB meta-bolic activity could extend the length of time that EBs maintain infectivity. Equalnumbers of purified EBs were incubated for 0, 18, and 24 h at 37°C in base medium (BM)or BM supplemented with various nutrients, including potential energy sources (Fig. 3).The BM consisted of IPB supplemented with 1% FBS, DTT, and a mixture of UTP, CTP,and GTP, to which we added a gradient of amino acids, ATP, or G6P. EB infectivity wasmeasured using reinfection via a traditional IFU assay, combined with automatedmicroscopy for enumeration of inclusions. Most bacteria can utilize multiple energysources, but it is not clear what range of metabolites the Chlamydia EB is capable ofutilizing. Therefore, we directly tested the ability of the EB to utilize amino acids, ATP,and G6P for maintenance of infectivity. We also tested the effect of the reversibleproton ionophore CCCP on EB infectivity. Incubation in BM resulted in �2-log reductionin EB infectivity over 24 h (Fig. 3). Dose-dependent maintenance of infectivity wasobserved upon supplementation of the base medium with G6P, molecular ATP, or amixture of amino acids, indicating that EBs can exploit a range of substrates to maintaininfectivity. The addition of CCCP to CIP-1 (BM plus 1 mM ATP plus 0.5 mM G6P plus 25mM amino acids) treatment led to a dramatic loss of infectivity, suggesting thatmembrane potential plays an important role in maintaining EB infectivity. These dataare consistent with other studies (11–13) suggesting that the EB is capable of metab-olism outside the chlamydial inclusion and that this metabolism may be necessary forthe EB to maintain an infectious phenotype. The combination of both ATP and G6P, aswell as the highest doses of ATP and G6P, did not substantially extend infectivitybeyond the level supported by substrates when supplemented separately, suggestingthat energy source availability is not sufficient for (indefinite) maintenance of infectivity(Fig. 3).

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G6P and amino acids support maintenance of EB ATP pools. Our infectivity datademonstrate that EBs can utilize at least three different energy sources to maintaininfectivity outside host cells. We previously demonstrated that G6P can serve as anenergy source for C. trachomatis EBs by supporting maintenance of EB ATP pools (13).To determine whether C. trachomatis EBs can also utilize amino acids for maintenanceof ATP pools, purified EBs were incubated in BM in the absence or presence of all 20amino acids (1.0 mM). After 6 h of incubation, the level of ATP extracted from bacteriaincubated in BM only was 1.5 � 0.2 nM, whereas the level extracted from bacteriaincubated in the presence of amino acids was 3.2 � 0.4 nM. When incubated in thepresence of amino acids and the protein synthesis inhibitor chloramphenicol (CAM), theextracted ATP levels were 4.0 � 0.7 nM, while the vehicle was only 3.25 � 0.4 nM (Fig.4A). Although only the CAM/amino acid treatment reached statistical significance, theoverall trend suggests that EBs can utilize amino acids to maintain ATP pools. Theincrease seen with CAM treatment suggests that by inhibiting protein synthesis, aminoacids are preferentially utilized for energy production (Fig. 4A). Incubation in theabsence of amino acids, but with G6P (100 �M), showed a similar albeit more robustresponse in EB ATP pools with extracted ATP levels of 5.6 � 0.5 nM (Fig. 4B). CAM

FIG 3 Nutrient availability enables EBs to maintain infectivity over an extended incubation period. The ability of EBs to actively maintain infectivity over 24 hin the presence of specific carbon and/or energy sources was tested by incubating purified EBs in axenic media, BM (CTP, GTP, and UTP at 1 mM, 1% FBS, and0.5 mM DTT), supplemented with G6P, amino acids, or ATP at different concentrations. The titer of infectious EBs at the end of the incubation period wasmeasured by enumeration of the IFU via a reinfection assay. Nonsupplemented BM resulted in a dramatic loss of infectivity by 18 h. Incubation in completeCIP-1 (BM plus 1 mM ATP plus 0.5 mM G6P plus 25 mM amino acids [AAs]) medium resulted in less than a 1-log reduction in infectivity over 24 h. The additionof the proton ionophore CCCP to CIP-1 medium resulted in a loss of infectivity. The addition of G6P at all concentrations enhanced the EBs’ maintenance ofinfectivity. A mixture of all 20 amino acids was also able to retard the loss of infectivity, and this effect was dose dependent. The addition of ATP also rescuedthe loss of infectivity in a dose-dependent manner. The experiment was run four times, and data from a representative experiment are shown. The depicteddata represent the means � the SEM (*, P � 0.01 [compared to BM alone]).

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treatment with G6P showed no significant changes in ATP maintenance (Fig. 4B).Overall, these data are consistent with the ability of EBs to maintain ATP levels viametabolism of a variety of nutrients, including amino acids.

De novo protein synthesis is not required for maintenance of infectivity.Previously published as well as current transcription data show that EBs in CIP-1 initiate denovo transcription and translation (13). Here, we have shown that the transcriptional profileof the EBs under these conditions is a mixture of expressed late chlamydial genes,suggesting an EB promoting profile, and a subset of noncanonically regulated genes. Ourdata also show that providing the EBs with diverse energy sources results in extendedinfectivity outside the cell. To determine whether de novo protein synthesis contributed tothe maintenance of infectivity, the effect of protein synthesis inhibition was tested (Fig. 5).EBs were incubated in complete CIP-1 in the presence or absence of CAM and rifampin (Rif),or CCCP. Inhibition of protein synthesis had no significant effect on maintenance ofinfectivity in the CIP-1 medium which contained both ATP and G6P, demonstrating thattranscription and protein synthesis were not required for long-term EB viability.

EBs undergo protein synthesis within the inclusion lumen. During the chlamyd-ial developmental cycle, EB production is asynchronous. EB production is initiated at�18 h postinfection and continues until cell lysis or inclusion extrusion at �48 hpostinfection (serovar L2). This produces a mixed population of EBs at the end of thecycle in that some EBs will have been present in the inclusion for 30 h, while others willonly have been in the inclusion for a few minutes. We hypothesized that the behaviorof EBs in CIP-1 may mimic the biology of the EBs inside the chlamydial inclusion. Todetermine whether the EB cell type undergoes maintenance metabolism until releasefrom the host cell, we evaluated the ability of intracellular EBs to undergo de novoprotein synthesis via incorporation of the methionine analog L-azidohomoalanine(AHA). AHA is incorporated during protein synthesis and can be detected fluorescentlyusing Click-iT chemistry (22). Cells were infected with C. trachomatis for 34 h and

FIG 4 Amino acid and G6P availability supports maintenance of ATP pools in purified EBs. The ability of purified EBs to generateATP from amino acids or G6P was tested. (A) EBs were incubated for 6 h in BM containing nucleotides other than ATP (1 mM), FBS(1%), and DTT (0.5 mM) supplemented with amino acids (1 mM). Although ATP levels in extracts from bacteria incubated innonsupplemented BM were depleted, bacteria incubated in the presence of amino acids had elevated ATP pools. Incubation inthe presence of the protein synthesis inhibitor CAM resulted in an increase in cellular ATP pools. (B) EBs incubated in the presenceof G6P (100 �M) resulted in CAM-independent maintenance of EB ATP pools. Depicted data were collected from at least twoindependent experiments and represent the means � the SEM.

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incubated with AHA for the final 2 h of a 36-h infection. To visualize all chlamydiae, thecells were labeled with an antibody raised against the major outer membrane protein(MOMP). AHA incorporation was analyzed by confocal microscopy. The images dem-onstrated that the majority of bacteria in the inclusion, EBs and RBs, were positive forAHA incorporation (Fig. 6). To verify that EBs were present in the inclusions at this timepoint, the ratio of IFU to GE was determined as described above. Infectious EBs werecalculated to be 78% � 2% of the total population of bacteria. As a control, prokaryoticprotein synthesis was inhibited with CAM, and no AHA incorporation was detected (Fig.6A). Notably, MOMP staining outside cells does not stain for AHA. It is difficult toevaluate these extracellular chlamydiae since it is unknown whether they were part ofthe inoculum that did not invade cells or are chlamydiae released from lytic events. Itis therefore impossible to know how long the EBs have been in the medium. Our datashow that maintenance of infectivity is not indefinite regardless of extracellular nutrientavailability; it is thus likely that the unlabeled bacteria are nonviable.

To verify that EBs were labeled by AHA, HeLa cells were infected with C. trachomatisfor 46 h and then pulsed with AHA for 2 h. The EBs were harvested using sonication andDG centrifugation (23, 24). Purified EBs were counterstained with DAPI (4=,6=-diamidino-2-phenylindole) to label cells containing DNA and visualized using confocal fluores-cence microscopy. The data showed that the majority of the DAPI-positive EBs werelabeled with AHA (Fig. 7A). In contrast, the CAM-treated control showed no AHAlabeling (Fig. 7A). Labeling of the EBs was also verified using flow cytometry. HeLa cellswere infected with C. trachomatis L2 transformed with a plasmid expressing thefluorescent protein Ruby2 under the control of the hctA promoter (see Fig. S5 in thesupplemental material). This construct expresses Ruby2 only late during infection,presumably only in EBs and transitioning RBs (see Fig. S5 in the supplemental material).Cells were infected for 46 h and then pulsed with AHA for 2 h. The EBs were harvestedand quantified using flow cytometry gated for Ruby2 and AHA incorporation (Fig. 7B).

FIG 5 Maintenance of infectivity in metabolically active extracellular EBs is independent of protein synthesis. To determine themetabolic basis for the maintenance of chlamydial infectivity, EBs were incubated in complete CIP-1 in the absence or presenceof specific inhibitors. The titer of infectious EBs at the end of the incubation period was measured by enumeration of IFU viaa reinfection assay. EBs were incubated with buffer in the absence of nutrients (BM), complete CIP-1, CIP-1 with CAM andrifampin (CAM�Rif), or CCCP. EBs maintained infectivity without active protein synthesis but required an intact membranepotential. The experiment was performed three times, and data from a representative experiment are shown. The depicteddata represent means � the SEM (*, P � 0.01) compared to IPB alone for CIP-1 and the CAM�Rif treatment. The CCCPtreatment was compared to CIP-1 treatment (*, P � 0.01). Overall, EB infectivity under axenic conditions appears to be largelydependent on energy metabolism.

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Quantification of the labeled cells revealed that 85% of the purified Ruby2-positive EBswere AHA labeled, while the CAM-treated cells showed only 6% of purified Ruby2-positive EBs labeled. Overall, these data demonstrate that EBs within the inclusionundergo active protein synthesis.

EBs maintain infectivity within the inclusion. EB metabolic activity under extra-cellular conditions significantly aids the maintenance of EB infectivity. Therefore, wehypothesized that the continued metabolic activity of EBs in the inclusion may play arole in maintaining EB viability and infectivity. To test the viability of interinclusion EBsover time, continued EB production after the first initial rounds was blocked bytreatment with penicillin. C. trachomatis does not use peptidoglycan as a structuralsacculus, and EBs do not contain a peptidoglycan cell wall. Instead, peptidoglycan aidscell septation by forming a ring at the cleavage furrow (25). Therefore, C. trachomatistreated with penicillin ceases to divide and does not produce new EBs (26). Themechanism linking the completion of cell division and EB development is not currentlyknown, but it has been reported in multiple studies that the addition of penicillin blocksEB formation (26–29). A recently published model of EB development also proposedcell division as being linked to EB development (30). To test the long-term viability ofEBs, C. trachomatis was allowed to develop for 36 h postinfection, at which point further

FIG 6 All cells in the chlamydial inclusion undergo active protein synthesis. The physiological state of C. trachomatis withinthe inclusion after RB differentiation was determined by assessing the overall protein synthesis by incorporation of themethionine analog azidohomoalanine (AHA) combined with Click-iT chemistry and confocal microscopy. Cos-7 cells wereinfected with C. trachomatis for 36 h and then pulsed with AHA for 2 h to label bacteria undergoing protein synthesis(green). (A) The AHA was labeled with Alexa Fluor 488. An anti-MOMP antibody was used to label Chlamydia (purple). Hostcells were labeled with DAPI to highlight nuclei. CAM was added to a subset of cultures at the time of AHA labeling to blockbacterial protein synthesis. (B) Magnified image of a chlamydial inclusion labeled with AHA-Alexa Fluor 488 and anti-MOMPantibody. The data illustrate that the entire C. trachomatis population appears to incorporate AHA and thus undergo activeprotein synthesis within the inclusion.

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EB production was inhibited by the addition of penicillin (Pen). Treated bacteria wereharvested at 48 (12 h Pen) and 60 h (24 h Pen) postinfection. Total bacterial cellnumbers were measured using quantitative PCR (qPCR) to calculate GE, and EB stabilitywas measured by a conventional IFU assay (Fig. 8). The IFU assay revealed that EBsproduced by 36 h were still infectious and viable 12 and 24 h after formation (Fig. 8A,compare 36 to 36 h � 12 h Pen and 36 h � 24 h Pen).

We showed that EBs in CIP-1 medium could utilize a variety of nutrients to maintaininfectivity through active metabolism but that, although the EBs were capable of

FIG 7 The vast majority of the EBs in the inclusion participated in protein synthesis during the AHA labeling period. EBs undergo proteinsynthesis within the inclusion. Host cells were infected with purified C. trachomatis and incubated for 46 h and then pulsed with AHA for2 h. (A) EBs were purified by sonication (to remove RBs) and density gradient centrifugation and then visualized by DNA staining with DAPIusing confocal microscopy. Nearly all DAPI-stained cells also showed incorporation of AHA. (B) Cos-7 cells were infected with C.trachomatis transformed with a plasmid expressing the fluorescent protein Ruby2 under the control of the hctA promoter. After 46 h, thecells were pulsed with AHA, the EBs were purified by density gradient centrifugation, and the AHA was labeled with Alexa Fluor 488. Thelabeled purified EBs were analyzed by flow cytometry. CAM was added at the time of AHA labeling to verify that incorporation was proteinsynthesis dependent. A gate was drawn around the AHA-positive cells to measure the percentages of bacteria that were labeled. A totalof 85% of the Ruby2-positive cells were positive for AHA, while after CAM treatment only 6% of the cells were AHA positive. The dataconclusively demonstrate that EBs continue to undergo protein synthesis even after terminal differentiation. Scale bar, 5 �m.

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detectable protein synthesis, de novo protein synthesis did not contribute to mainte-nance of EB infectivity. To determine whether EB protein synthesis was important formaintaining infectivity in the inclusion, infected cultures were treated with CAM inaddition to penicillin for the indicated times. Again, EBs were allowed to develop for 36 hprior to Pen and CAM treatment and harvested at 48 and 60 h postinfection, and both GEand infectious progeny were determined. Inhibition of protein synthesis with CAM,which completely inhibited AHA incorporation (Fig. 6 and 7), had no significant effecton long-term EB infectivity (Fig. 8A). As has been reported previously (28), Pen treat-ment alone did not inhibit genome replication, but in combination with CAM genomereplication was arrested. This suggests that the axenic conditions are mimickingconditions relevant for the inclusion lumen and that the EB likely relies on energymetabolism but not protein synthesis both outside cells and in the inclusion tomaintain infectivity. Indeed, CIP-1 was designed with the concentrations of Na� and K�

according to levels measured for the inclusion lumen (31) consistent with the ionicconditions of CIP-1 reflecting the inclusion environment.

DISCUSSION

Functional specialization between EB and RB cell forms is a defining characteristic ofC. trachomatis, and completion of the pathogen’s biphasic developmental program isessential for infectivity, viability, and virulence. The EB cell type is wholly responsible forinitiating new rounds of infection, mediating cell entry and initiating the creation of theintracellular replication niche, while the RB cell type replicates and initiates differenti-ation to the EB cell type to prepare for release and reinfection. The production of theEB cell type is asynchronous, leading to EBs of vastly different ages present in theinclusion until released by host cell lysis or extrusion of the inclusion (32). The releaseof chlamydial EBs at the end of the intracellular developmental cycle by the aforemen-

FIG 8 EBs generated early in infection maintained infectivity through the developmental cycle. The viability of EBs made early duringinfection was determined by using a reinfection assay to quantify infectious EBs over time after new EB production was inhibited withpenicillin (Pen). Cells infected with C. trachomatis for 36 h were treated with Pen to inhibit further development of EBs and incubated foran additional 12 and 24 h. EB infectivity was quantified using a reinfection assay (A), and GE were calculated using qPCR (B). The additionof the protein synthesis inhibitor CAM did not significantly affect the data obtained using Pen alone. No difference was seen between the36-h time point and either Pen only or Pen plus CAM treatment at plus 12 h, and only a slight reduction in infectivity was seen at plus24 h, again with no difference between Pen and Pen plus CAM at either time point. The experiment was repeated three times, and datafrom a representative experiment are shown. The depicted data represent the means � the SEM.

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tioned mechanisms has been the focus of research for several decades. Recentlypublished data showing that the chlamydial EB can metabolize extracellularly (11),along with data showing that the chlamydial EB can respond to specific energy sources(12, 13), suggest that the simple model that the EB is metabolically inert after formationand is a passive entity waiting for contact with the next host cell may be an oversim-plification of its role in pathogenesis. The data presented here illustrate that EBs canremain infectious for at least 24 h and utilize multiple energy sources to maintaininfectivity. We propose a model whereby the EB actively maintains infectivity in at leastthree distinct environments and phases of the chlamydial developmental cycle, namely,upon deposition at mucosal surfaces, within the inclusion after terminal RB-EB devel-opment, and within extrusions (Fig. 9).

Distinct temporal gene expression in Chlamydia controls the phenotypic separationbetween the RB and EB cell types. By assessing the transcriptome of EBs incubated inthe presence of different nutrients, we found evidence suggesting that a gene expres-sion profile is maintained that most closely resembles isolated EBs. This is consistentwith our data showing that under extracellular conditions, EBs actively maintain the

FIG 9 Model of EB maintenance cycle. The chlamydial developmental cycle has historically been described andcharacterized from the time of host infection by an EB to the time of terminal RB-EB differentiation in the lateinclusion. The chlamydial EB itself is typically presented as a largely dormant infectious particle in this develop-mental cycle. Recent independent studies, along with data presented here, support a far more complex picture inwhich the EB itself can be viewed as having a “maintenance cycle.” During the extracellular-maintenance phase, EBsdeposited onto mucosal surfaces actively maintain infectivity by metabolizing nutrients released by the mucosalepithelium as a means to enhance virulence. During the intracellular-maintenance phase, EBs within the interin-clusion space (inclusoplasm) have access to nutrients and remain metabolically active as evidenced by continuousprotein synthesis. EBs maintained within extruded inclusions are also expected to continue active metabolism sincethis hybrid environment may have a favorable nutrient makeup. Infection requires active type III secretion (green)and reorganization of the host actin cytoskeleton (red), and the activity of this system is known to be ATPdependent. The maintenance of a functional type III secretion system and secretion of its early effectors arepotentially the critical system dependent on ATP pools maintained through active metabolism.

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infectious EB phenotype. Comparison of the EB transcript profile after incubation underspecific nutrients or during early intracellular development suggests that EB germina-tion is regulated by a transcriptional program that is not recapitulated under the hostcell-free conditions tested in this study. For example, crucial to germination is therelease of the chlamydial histones, HctA and HctB, from the chromosome immediatelyupon cell entry. This process is facilitated by the IspE enzyme of the isoprenoidsynthesis pathway (14, 33). Our data show ispE was induced 3.5-fold (P � 0.01) at 2 hand 3.8-fold (P � 0.01) at 5 h but was not induced at all under the extracellularincubation conditions. As illustrated in other bacterial systems, a specific physiologicalstate (e.g., replication) can present with diverse gene expression patterns (34). BecauseChlamydia has a reduced genome and a developmental cycle that is specially adapted,this is less likely to be true. There were a significant number of genes induced in theextracellular media that did not conform to canonical “late” EB-like or germination“early” genes as defined by Belland et al. (8) and Shaw et al. (17). These genes includedtranscripts for amino acid production and import, inclusion membrane proteins, thetype II secretion operon and, surprisingly, the early transcriptional control protein Euo.Euo expression during incubation in CIP-1 (G6P, 0.5 mM) was only modestly induced(1.5-fold, P � 0.01) compared to the EB inoculum. The significance of this induction isnot clear since genes reported to be repressed by Euo (35) had a variable response. Forinstance, the late gene ltuB was induced 3.7-fold, and hctB was induced 5.1-fold (P �

0.01) in CIP-1 (G6P 0.5 mM), while pgk and bioY were downregulated 2.0- and 1.2-fold,respectively (P � 0.01). It is uncertain whether the observed expression profile of thesegenes is an artifact of the medium or an active response to reflect gene expressionunder conditions where the EB is simply maintaining viability and infectivity. Becausenutritional and physicochemical conditions in extracellular environments encounteredby EBs (e.g., mucosal surface) are expected to be more diverse than an intracellular (i.e.,inclusion) environment, the gene expression profile of extracellular EBs undergoingmaintenance metabolism may show significant variation.

EBs do not synthesize considerable levels of protein de novo in medium containingamino acids only but did with access to G6P (13). In this study, inhibition of proteinsynthesis resulted in no loss of EB infectivity over 24 h intracellularly and extracellularly.Therefore, gross de novo protein synthesis appears to have little role in maintenance ofEB infectivity. On the other hand, disruption of the EB membrane potential by thereversible proton ionophore CCCP resulted in near complete loss of infectivity. There-fore, we conclude that maintenance of EB infectivity appears to be largely dependenton the ability of EBs to maintain cellular energetics, including a membrane potential.The significance of EB protein synthesis after development is uncertain but may reflectcurrently undescribed EB functions or may reflect residual translation that is detectedby the very sensitive AHA Click-iT and [35S]cysteine-methionine labeling assays. Main-tenance of chlamydial infectivity within both the inclusion and extracellular environ-ments has only received minimal attention, likely in part due to the long-held view thatthe chlamydial EB is metabolically inert. Indeed, the more recently introduced conceptof the extrusion of intact inclusions (32, 36), in addition to the release of EBs from thehost by lysis, has been presented as a potential mechanism for chlamydiae to survivefor longer periods of time following release from the host cell.

Promiscuity in the EB’s ability to use a wide range of substrates to maintaininfectivity is consistent with evolution of a cell form specialized to initiate new infectionevents under conditions of varying nutrient availability. In our studies, EB maintenanceof infectivity was dependent on any one of three potential energy sources outside thecell: G6P, ATP, or amino acids. Chlamydia can directly use ATP as an energy source byimporting it from the cell through ADP/ATP exchange (37, 38). Likewise, C. trachomatisencodes a G6P transporter (UhpC) and metabolic pathways to undergo oxidative andsubstrate-level production of ATP from G6P (39). However, the ability of C. trachomatisto generate energy from amino acids is less well understood. In this study, we providedall 20 amino acids and therefore cannot conclude which amino acid(s) was utilized formaintenance of infectivity in the EB. However, measurement of cellular ATP pools

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following incubation of purified EBs with amino acids demonstrates that amino acidssupport the maintenance of EB ATP pools (Fig. 4). C. trachomatis encodes a number ofamino acid transporters, but it is unclear whether all of them function during the EBdevelopmental stage (39). In our experiments, the methionine analog AHA was effi-ciently taken up by EBs in the inclusion; thus, it is likely that at least the BrnQbranched-chain amino acid permease is functional in the EB. BrnQ transports valine,leucine, isoleucine, methionine, and phenylalanine (40). In addition, it has been re-ported that phenylalanine is taken up by the EB (11). Of these amino acids, valine,leucine, methionine, and phenylalanine could potentially act as substrates for thetruncated C. trachomatis tricarboxylic acid (TCA) cycle (39, 41, 42). Recent studiesinvolving the environmental organism Protochlamydia suggest that amino acids are thepreferential carbon source during early stages of development with a postreplicativeswitch to glucose metabolism (43). It is clear that understanding the metabolism of thechlamydial cell forms will lead to greater understanding of the complex developmentalcycle that underlies its pathogenesis.

In cultured cells ATP, amino acids and G6P pools are unlikely to be limiting. ATPlevels in cultured cells have been measured to be ca. 4 to 5 mM (44), the glucoseconcentration in RPMI cell culture medium is 11 mM, and each amino acid is presentat �1 mM. These measurements do not take into account the nutrients available fromserum. However, during infection, one of the major effectors of the interferon gammaresponse in human cells is the induction of the enzyme indoleamine 2,3-dioxygenase(IDO). IDO catalyzes degradation of the essential amino acid tryptophan (45). Theinduction of IDO and depletion of tryptophan has been shown to mediate interferongamma-induced inhibition of intracellular Chlamydia replication (46). Intriguingly, tryp-tophan is unlikely to be utilized by the chlamydial TCA cycle as an energy substrate (39,41, 42), and we have shown that protein synthesis has no effect on EB maintenance ofinfectivity, suggesting that the EB, once produced, may be immune to the inhibitoryeffects of tryptophan starvation.

The role of the EB cell type in chlamydial pathogenesis has been thought to belimited to the passive action of initiating the next round of infection through chanceencounter with a susceptible host cell immediately after release from the host cell bylysis. It is intriguing that the EB is metabolically active with a promiscuous range ofsubstrate utilization, potentially allowing the EB to respond to diverse environmentalniches (Fig. 9). We show here that the EB can survive inside the inclusion during aproductive infection for at least 24 h after differentiation, but it is an interestingquestion as to how long the EB can maintain metabolism and the infectious state.Under stress conditions, C. trachomatis responds by forming aberrant/persistent cellforms and no longer generates EBs (47). Inclusions in this state are thought to persistfor months (48). It is possible that the aberrant inclusion remains a favorable environ-ment for the EBs already present to maintain infectivity.

The EB is also subjected to other environmental niches outside the host cell,including direct contact with extracellular mucosal secretions and the chlamydialexosome. Compounds secreted by epithelial cells and the accumulation of metabolitesreleased from dying epithelial cells creates a complex nutritional environment atmucosal surfaces. Specific metabolites detected in mucosal liquid include amino acids,pyruvate, glucose, phospholipids, and nucleotides, including ATP (49–51). Of potentialsignificance, C. trachomatis has been shown to stimulate release of ATP from infectedHeLa cells (52). Pathogen-dependent release of ATP at mucosal surfaces could prolongEB infectivity increasing the efficiency of infection of neighboring host cells. Theexosome could be considered a hybrid environment maintaining some of the environ-mental conditions of the intracellular inclusion in combination with access to theextracellular environment. Extrusion of the chlamydial inclusion from cells occurs inroughly half of infected cells in culture, and the extruded chlamydial exosomes havebeen detected during in vivo infections of mice (32). The environmental and nutritionalconditions within these exosomes has not yet been described, but it is conceivable thatthe exosome represents a nutritionally favorable environment for the maintenance of

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EB infectivity to increase the transmission and spread of the infection. The reduction ofdisulfide bonds in the outer membrane of chlamydial EBs has been postulated toenhance nutrient transport (53). Although not required for EB metabolic activity, weused the reducing agent DTT in the present study to facilitate EB activity (13). The effectof mucosal antioxidants such as glutathione on EB activity and the maintenance ofinfectivity remains to be tested.

Overall, this study supports the idea that the chlamydial EB is an actively maintainedcell form that responds to nutrients and environmental signals to maintain infectivityin multiple environments, thus aiding in chlamydial pathogenesis. This report alsoindicates that stimulation of metabolism by chemically diverse substrates does nottrigger a gene expression pattern suggestive of differentiation. Identification of condi-tions or molecular triggers of the transcriptional program that governs EB-RB differen-tiation is a critical question in chlamydial biology.

MATERIALS AND METHODSOrganisms and cell culture. C. trachomatis serovar L2 (LGV Bu434) was grown in McCoy cells.

Reticulate bodies (RBs) and elementary bodies (EBs) were purified by DG centrifugation essentially asdescribed previously (23) following 17 h or 43 to 45 h of infection, respectively. EBs were stored at �80°Cin sucrose-phosphate-glutamate buffer (10 mM sodium phosphate [8 mM K2HPO4, 2 mM KH2PO4], 220mM sucrose, 0.50 mM L-glutamic acid; pH 7.4) until use (54). Transmission electron microscopy wasused to confirm EB and RB ultrastructure, and inclusion-forming unit (IFU) assays were used incombination with qPCR to determine the ratio of infectious particles to GE as a measure of stockpurity and quality (GE/IFU ratio approaching 1:1). Cell lines were obtained from American TypeCulture Collection. Cells were maintained in RPMI 1640 medium (Cellgro) supplemented with 10%FBS and 10 �g/ml gentamicin.

Infection of cultured cells for RNA-seq analysis. Confluent monolayers of host cells were incu-bated with C. trachomatis EBs for 10 min in HBSS at room temperature while rocking at an MOI of �1,000for 2- and 5-h infections and at an MOI of �5 for 12, 18, and 24 h (Gibco). For the 2- and 5-h infectionsthe inoculum was removed, and the cultures were washed five times with HBSS containing 100 �g/mlheparin. For the 12-, 18-, and 24-h time points the cells were incubated with the inoculum for 30 minbefore washing. The final wash was replaced with fresh complete medium, and the infection was allowedto continue for the reported incubation times.

Axenic incubation for analysis of infectivity. For analysis of maintenance of infectivity, EBs wereincubated in six-well plates under the indicated nutrient conditions at 109 GE/ml using 3 ml of mediumper well. Duplicate samples were incubated in a CO2 incubator at 37°C. At the indicated time points, thewell contents were mixed, 100 �l was removed and combined with 900 �l of K-36 buffer, and thesamples were then stored at �80°C until analysis by IFU assay.

Immunofluorescence staining of cultured cells. Cells for fluorescence microscopy were grown on12-mm number 1.5 borosilicate glass coverslips coated with poly-L-lysine (Sigma). Samples were fixed in4% paraformaldehyde. To visualize primary antibody binding, appropriate Alexa Fluor (Molecular Probes/Life Technologies)-conjugated secondary antibodies were used: Alexa Fluor 488/568/647 conjugated toantibodies against mouse or human immunoglobulin G (IgG).

Microscopy. Fluorescence images were acquired using a Nikon spinning disk confocal system witha 60� oil immersion objective lens, equipped with an Andor Ixon EMCCD camera, under the control ofthe Nikon elements software. Images were processed using the image analysis software ImageJ (http://rsb.info.nih.gov/ij/). Representative confocal micrographs displayed in the figures are maximal intensityprojections of the three-dimensional data sets, unless otherwise noted. ImageJ was also used to quantifyfluorescent signal for measuring inclusion intracellular trafficking. Live cell imaging of inclusions express-ing the fluorescent protein Ruby2 (55) under the control of the hctA promoter (56) was achieved usingan automated Nikon epifluorescence microscope equipped with an Okolab heated stage and an AndorZyla sCMOS camera. Images were taken every 15 min for 48 h. Multiple fields of view of multiple wellsof a glass-bottom 24-well plate were imaged. The fluorescence intensity of each inclusion over timewas tracked using the ImageJ plug-in TrackMate (57), and the results were averaged and plottedusing Python and matplotlib (see Fig. S5 in the supplemental material). For analysis of the RB andEB ultrastructure, bacteria were purified from cultured McCoy cells following 17 h or 43 to 45 h ofincubation as described, respectively. Bacteria were washed in K-36 buffer (0.1 mM potassiumphosphate buffer [pH 7.0], 0.1 mM potassium chloride, 0.015 mM sodium chloride) and thenresuspended and fixed (2% paraformaldehyde, 2% glutaraldehyde, 0.1 M cacodylate buffer; pH 7.2).After fixation, the samples were rinsed and dehydrated before embedding with Spurr’s resin.Ultrathin sections (70 to 100 nm) were prepared with an ultramicrotome (Reichert Ultracut R; Leica)and placed on Formvar-coated slot grids. Samples were stained with 2% uranyl acetate andpoststained with Reynolds lead citrate. The sections were imaged using a FEI Tecnai G2 transmissionelectron microscope (FEI Company) (see Fig. S2 in the supplemental material). Changes to contrastand signal intensity were applied to the entire image.

Sequencing. Total RNA was isolated from both infected cells and treated purified EBs, in quadru-plicate, using TRIzol reagent (Life Technologies) according to the protocol provided. RNA samplesisolated from infected cells were further enriched by the removal of eukaryotic and bacterial rRNA

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(Ribo-Zero Gold rRNA kit; Illumina, Inc.), eukaryotic mRNA [Dynabeads Oligo(dT)25; Life Technologies],and genomic DNA (Turbo DNA-free kit; Invitrogen). RNA isolated from purified EB samples were furtherenriched by the removal of bacterial rRNA (Ribo-Zero GramNegative bacteria kit; Illumina, Inc.) andgenomic DNA (Turbo DNA-free kit). The enriched RNA samples were quantified, and the libraries werebuilt and barcoded by the IBEST Genomics Resources Core at the University of Idaho. The libraries weresequenced by the Vincent J. Coates Genomics Sequencing Laboratory at UC Berkeley on a HiSeq2500 50SRR platform. RNA-seq reads were aligned to the published C. trachomatis L2 Bu434 genome using thebowtie2 aligner software (58). Reads were quantified using HTseq (59). Statistical analysis and normal-ization of read counts was accomplished using DESeq2 in R (19). Log2 fold changes and statistics werealso calculated using DESeq2. Heatmaps and hierarchical clustering were generated and visualized usingPython with Pandas and the Seaborn visualization package (60). Aligned reads are accessible from theNCBI’s Sequence Read Archive (SRA), submission number SUB2915657.

Protein labeling. Chlamydial specific protein synthesis was detected using the Click-iT AHA AlexaFluor 488 protein synthesis HCS assay (Invitrogen). Cos-7 cells were grown on 12-mm number 1.5borosilicate glass coverslips coated with poly-L-lysine (Sigma) and infected with C. trachomatis serovar L2(LGV Bu434) at an MOI of �1. Cos-7 cells were chosen for this assay since eukaryotic protein synthesiswas completely inhibited in this cell line. Infected cells were starved, and eukaryotic protein synthesiswas inhibited 3.5 h prior to the time point of interest for 30 min in amino acid labeling medium (RPMI1640 medium without L-glutamine, L-cysteine, L-cystine, or L-methionine [MP Biomedicals], supple-mented with 10% Fetalplex serum [Gemini Bio Products], 2 mM L-glutamine plus 10 �g/ml emetine, and100 �g/ml cycloheximide). Starved infected cells were then incubated for a further 3 h in complete�Metmedium supplemented with 100 �M AHA, at which point samples were fixed in 4% paraformaldehyde.Click-iT chemistry was performed as described by the kit protocol with the following minor additions toidentify C. trachomatis. After the Click-iT reaction, samples were washed once in 3% bovine serumalbumin, followed by incubation at 4°C overnight with a 1:100 dilution of the MOMP monoclonalantibody clone 142H (Thermo Scientific). Alexa Fluor 647-conjugated secondary antibodies againstmouse IgG were used (Molecular Probes/Invitrogen) to detect the primary antibody. The far-redfluorescent DNA dye DRAQ5 (Biostatus) was used to visualize nuclei.

To visualize protein synthesis in purified EBs, Cos-7 cells were infected with C. trachomatis at an MOI of�1 and treated as described above 3.5 h prior to density gradient purification of the EBs at 48 h postinfection.The final EB pellet was subjected to Click-iT to label the incorporated AHA as described above.

Measurement of ATP pools. Bacterial ATP pools were measured using a luciferase-based ATPdetermination kit (Molecular Probes). Briefly, EBs were incubated under the indicated conditions in 1-mlmicrocentrifuge tubes using 500-�l volumes and 109 EBs per sample for 6 h at 37°C. CAM was dissolvedin dimethyl sulfoxide (DMSO) and used at a final concentration of 10 �g/ml, diluting the DMSO solvent1:2,000. After incubation, the samples were cooled on ice, and the bacteria were pelleted by centrifu-gation (4 min, 20,000 � g) at 4°C and then washed in 100 �l of ice-cold K-36 buffer. Pellets wereresuspended in 10 �l of 0.01% (wt/vol) sodium dodecyl sulfate (SDS) in distilled H2O, heated to 95°C for5 min to extract ATP, and mixed with 90 �l of phosphate-buffered saline. The concentration of ATP wasmeasured against an ATP standard curve in the presence of 0.001% SDS.

IFU determination. EBs were released from infected cells using sonication and pelleted by centrif-ugation for 30 min at 14,000 rpm. Similarly, EBs treated in axenic media were pelleted by centrifugationfor 30 min at 14,000 rpm. The EB pellets were resuspended in HBSS. Cos-7 cells were plated into 96-wellclear plates the day before. Infections were carried out by performing 2-fold dilutions of the EBs andincubated for 30 min at 37°C. After the incubation, the HBSS was replaced with complete mediumcontaining 1 �g/ml cycloheximide. The plates were incubated for 30 h before fixing in methanol. EBswere stained with a fluorescein isothiocyanate-labeled anti-MOMP polyclonal antibody (Thermo Fisher,catalog no. PA1-73073), and cell nuclei were stained with DAPI. Nine images per well over four dilutionswere captured using an automated script with a Nikon TE2000 inverted microscope under the control ofthe �Manager software equipped with a computer-controlled XYZ stage (61, 62). Inclusion numbers weredetermined using a custom analysis script in the Fiji software (63, 64). Inclusions were detected by settingan empirically determined threshold and object detection. Statistical analysis and visualization of the rawinclusion counts were carried using custom Python scripts with the Python pandas data analysis package(http://pandas.pydata.org).

Determination of genome equivalents. Genomic DNA was isolated from either purified EBs orinfected cells using an Invitrogen genomic mini-DNA purification kit. Genome copy numbers weredetermined using hctA-specific primers and SYBR green detection using a StepOnePlus real-time PCRsystem (Thermo Fisher Scientific).

Flow cytometry. EBs expressing the Ruby2 protein under the control of the hctA promoter werelabeled for 2 h with AHA at 46 h postinfection and purified as described above. The incorporated AHAwas labeled using Click-iT as described above. The fluorescent signals in these purified labeled EBs werequantified using a Beckman Coulter CytoFLEX S flow cytometer to collect the signal in both the mCherrychannel and the Alexa Fluor 488-labeled AHA channel. Visualization and statistical analysis of thefluorescent signals for each cell was performed using Python with pandas and the Flow Cytometry Toolslibrary (http://eyurtsev.github.io/FlowCytometryTools/).

Statistical analysis. For analysis of the IFUs, all experiments were performed in duplicate with atleast two experimental replicates. Statistical significance was determined using Student t test for pairwisecomparisons and analysis of variance for comparisons of more than two groups.

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SUPPLEMENTAL MATERIAL

Supplemental material for this article may be found at https://doi.org/10.1128/JB.00065-18.

SUPPLEMENTAL FILE 1, PDF file, 2.5 MB.SUPPLEMENTAL FILE 2, XLSX file, 1.5 MB.SUPPLEMENTAL FILE 3, XLSX file, 0.1 MB.

ACKNOWLEDGMENTSWe thank Viveka Vadyvaloo for critical review of the manuscript. We thank Dan

Mullendore, Franceschi Microscopy and Imaging Center at Washington State Universityfor technical assistance with electron microscopy. The Vincent J. Coates GenomicsSequencing Laboratory at UC Berkeley, supported by NIH S10 instrumentation grantsS10RR029668 and S10RR027303, was used for sequencing analysis.

This study was supported by 1R21AI113617-01 (S.G., N.G., and A.O.), 1R21AI115244-01(A.O.), and laboratory start-up funds from Washington State University (A.O.) and theUniversity of Idaho (S.G.).

Author contributions were as follows: conceptualization, S.G., N.G., T.H., and A.O.;data curation, S.G., N.G., H.Y., B.B., and A.O.; formal analysis, S.G., N.G., H.Y., B.B., T.H., andA.O.; funding acquisition, S.G., N.G., and A.O.; investigation, S.G., N.G., and A.O.; projectadministration, S.G. and A.O.; resources, S.G., N.G., and A.O.; supervision, S.G., N.G., andA.O.; visualization, S.G., N.G., and A.O.; and manuscript preparation, S.G., N.G., and A.O.

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