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Actin polymerization in the cytosol and at the plasma membrane is locally regulated by actin nucleators. Several microbial pathogens exploit cellular actin polymerization to spread through tissue. The movement of the enteric pathogen Shigella flexneri, both within the cell body and from cell to cell, depends on actin polymerization. During intercellular spread, actin polymerization at the bacterial surface generates protrusions of the plasma membrane, which are engulfed by adjacent cells. In the cell body, polymerization of actin by Shigella spp. is dependent on N-WASP activation of the Arp2/Arp3 complex. Here we demonstrate that, in contrast, efficient protrusion formation and intercellular spread depend on actin polymerization that involves activation of the Diaphanous formin Dia. While the Shigella virulence protein IpgB2 can bind and activate Dia1 (N. M. Alto et al., Cell 124:133-145, 2006), its absence does not result in a detectable defect in Dia-dependent protrusion formation or spread. The dependence on the activation of Dia during S. flexneri infection contrasts with the inhibition of this pathway observed during vaccinia virus infection.
During infection, several human bacterial pathogens enter host cells and spread through host tissues by moving directly from one cell into adjacent cells. These microorganisms, including Shigella spp., Listeria monocytogenes (44), Rickettsia spp. (43), Burkholderia spp. (21), and Mycobacterium marinum (42), induce the polymerization of host actin into tails that propel them through the cell cytoplasm to the cell periphery. Actin tail assembly in the cell body involves local activation of actin polymerization through the Arp2/Arp3 (Arp2/3) complex (6, 11, 14, 19, 27, 49). The Arp2/3 complex initiates new filament assembly and cross-links those filaments at 70° angles (28). At the cell periphery, Shigella spp. push outwardly against the plasma membrane, creating a membrane-bound cell extension (“protrusion”) that extends tens of micrometers from the cell surface and contains a bacterium at its tip (5). Contact of a protrusion tip with the membrane of an adjacent cell is followed by its uptake into the adjacent cell by a process that resembles macropinocytosis (20), leading to the spread of the infection into adjacent cells.
Although it is clear that actin assembly is required for the formation of protrusions by Shigella spp., the specific molecular mechanisms involved are poorly understood. Shigella spp. frequently form protrusions in tissue culture cells at sites of focal adhesions (30). The actin network at the base of protrusions contains filaments that are oriented in parallel arrays, in contrast to the angled arrays of actin filaments that predominate in actin tails associated with bacteria in the cell body (15), suggesting that actin nucleation processes independent of the Arp2/3 complex may be involved in protrusion formation.
Formins are ubiquitously expressed proteins that, like the Arp2/3 complex, initiate de novo polymerization of actin (31, 36). In contrast to Arp2/3 complex-mediated actin polymerization, formin-mediated actin polymerization leads to cross-linking of actin polymers in parallel arrays (31, 36). Formins play critical roles in a variety of cytoskeletal processes in different cell types, including cytokinesis, cell polarity, cell migration and adhesion, and intracellular trafficking (13). At the cell membrane, the mammalian Diaphanous-related formins Dia1 and Dia2 function as effectors of the small GTPase RhoA (1, 39, 48). RhoA plays a critical role in the generation of actin stress fibers that attach at adherens junctions and focal adhesions. The localization of Dia1 and Dia2 at sites of potential Shigella flexneri exit from the cell makes them ideal candidates as mediators of protrusion formation. Their potential role in this process is examined here.
The wild-type S. flexneri strain used in this study is serotype 2a strain 2457T (23). The conditional virB mutant, 2457T virB::Tn5/pDSW206-Ptsc-virB, has been described previously (25). An isogenic ipgB2 mutant was generated by deleting the coding sequence of ipgB2 and inserting a kanamycin cassette via phage λ Red recombinase-mediated homologous recombination (10). Following P1-mediated transduction of the kanamycin-resistant locus into a clean 2457T background, the kanamycin cassette was removed using FLP recombinase to generate a nonpolar, unmarked isogenic ipgB2 mutant (10). The lack of the ipgB2 coding sequence, the lack of the kanamycin cassette, and the maintenance of the flanking DNA sequences were verified by PCR. Bacteria were grown in tryptic soy broth from individual colonies that were red on agar containing Congo red.
pCMV-Myc (carrying Myc), pDsRed-Monomer-N1 (carrying DsRed), and pEGFP-C1, pEGFP-C2, and pEGFP-N1 (carrying enhanced green fluorescent protein [EGFP]) were obtained from Clontech. pEGFP-C1-IpgB2 was created via the site-specific Gateway (Invitrogen) recombination system. pEGFP-C1-Dia1 (16), encoding murine Dia1, was a gift from Naoki Watanabe. pEGFP-N1-Dia1129-369 (DID-EGFP), encoding the murine Dia1 Diaphanous inhibitory domain (DID), and pEGFP-N1-Dia1129-369(A256D) [DID(A256D)-EGFP], encoding the Diaphanous autoregulatory domain (DAD)-binding mutant of the murine Dia1 DID, EGFP-DID, and EGFP-DID(A256D) were gifts from Henry N. Higgs. Murine and human Dia1 sequences are 86 to 88% identical, and murine and human Dia2 sequences are 84% identical. pMyc-RhoA(T19N) [encoding Myc-RhoA(T19N)] was a gift form Ralph R. Isberg; pGFPmut2 (8) was a gift from Brendan Cormack; and pSUPER-retro, pSUPER-retro-mDia1KD1, and pSUPER-retro-mDia1KD2, for interfering RNA (RNAi) for Dia1 (35), were gifts from Leonidas Tsiokas. For experiments to assess protrusion formation following depletion of Dia1 or Dia2, a mixture of pSUPER-retro-mDia1KD1 and pSUPER-retro-mDia1KD2 or Dharmacon siGENOME small interfering RNA (siRNA) D-010347-01 was used to target Dia1, while V2HS_73202 short hairpin RNA (shRNA) (OpenBiosystems) or Dharmacon siGENOME siRNA D-018997-02-0002 was used to target Dia2. For plaque assays, RNA interference with Dia1 was performed using Dharmacon SMARTpool siRNA M-010347-02, with an siRNA that targets gfp mRNA as a control. pDsRed-Monomer-N1-Dia1129-369 (DID-DsRed) and pDsRed-Monomer-N1-Dia1129-369(A256D) [DID(A256D)-DsRed] were generated by cloning the XhoI-EcoRI fragments from DID-EGFP and DID(A256D)-EGFP, respectively, into pDsRed-Monomer-N1. pCMV-Myc-Dia1129-369 (Myc-DID) and pCMV-Myc-Dia1129-369(A256D) [Myc-DID(A256D)] were generated by cloning PCR-amplified DNA encoding the indicated residues from EGFP-DID and EGFP-DID(A256D), respectively, into the SalI and NotI sites of pCMV-Myc. pCMV-Myc-Dia1 (carrying Myc-Dia1) was generated by cloning a PCR-amplified Myc sequence into pEGFP-C1-Dia1. pCMV-Myc-IpgB2 (carrying Myc-IpgB2) was generated by cloning PCR-amplified DNA encoding the full coding sequence of IpgB2 into the EcoRI and NotI sites of pCMV-Myc. The sequences of primers used in PCR and sequencing are available from the authors upon request.
PtK2 rat kangaroo kidney epithelial cells were maintained in Dulbecco's modified Eagle's essential medium (DMEM), supplemented with 0.1% glucose and 10% fetal bovine serum, under humidified air containing 5% CO2 at 37°C. HeLa cells were maintained under the same conditions in minimal essential medium (MEM) supplemented with 10% fetal bovine serum. For analysis of the induction of formation of stress fibers by IpgB2, 3 × 106 HeLa cells were transfected by electroporation with EGFP and either Myc-IpgB2 or Myc at a ratio of 1:5 using a total of 6 μg of DNA in 300 μl serum-free MEM. Electroporations were performed using a Bio-Rad Gene Pulser II electroporation system with a 4-mm-diameter cuvette at 0.250 kV and 950 μF. A 50-μl portion from each transfection was seeded onto acetone-rinsed coverslips in 2 ml of medium and was incubated overnight at 37°C. Sixteen hours posttransfection, cells were fixed with 3.7% p-formaldehyde in cytoskeleton fix buffer [F buffer, comprising 5 mM KCl, 137 mM NaCl, 4 mM NaHCO3, 1.1 mM Na2HPO4, 0.4 mM KH2PO4, 2 mM MgCl2, 5 mM piperazine-N,N′-bis(2-ethanesulfonic acid) (PIPES), 2 mM EGTA, and 5.5 mM glucose (pH 7.2)] and were permeabilized with 0.5% Triton X-100 in F buffer. Polymerized actin was labeled with Alexa Fluor 568 phalloidin (Invitrogen). Transfected cells were identified by green fluorescence and were scored for increased stress fiber formation relative to that of control cells. The results for PtK2 and HeLa cells were similar.
For analysis of the colocalization of IpgB2 and Dia1, PtK2 cells were transfected by electroporation with GFP-IpgB2 and either Myc-Dia1 or Myc at a ratio of 1:1, as described above. Sixteen hours posttransfection, cells were fixed and permeabilized as described above. Labeling of the Myc tag was performed using a monoclonal anti-Myc antibody (Clontech) and a Texas red-conjugated anti-mouse secondary antibody (Jackson ImmunoResearch).
For analysis of the inhibition of protrusion formation by the DID, PtK2 cells were transfected with EGFP, DID-EGFP, or DID(A256D)-EGFP and were seeded as described above. Sixteen hours posttransfection, cells were infected with exponential-phase wild-type S. flexneri at a multiplicity of infection (MOI) (ratio of bacteria to cells) of 20 at 37°C, as described previously (2). Following an initial invasion period of 1.5 h, cells were washed, and the infection was allowed to continue for an additional 1.5 h in the presence of 50 μg/ml gentamicin, which kills extracellular but not intracellular bacteria. Cells were fixed with 3.7% p-formaldehyde in F buffer and were permeabilized with 0.5% Triton X-100 in F buffer. Polymerized actin was labeled with Alexa Fluor 568 phalloidin (Invitrogen), and DNA was labeled with 4′,6-diamidino-2-phenylindole (DAPI; Invitrogen). Transfected cells were identified by green fluorescence. Images of transfected cells that were infected were acquired and were analyzed for the total number of intracellular bacteria, the number of intracellular bacteria with actin tails, the number of intracellular bacteria in protrusions, and the lengths of actin tails. A protrusion was defined as an extension of the plasma membrane outside the normal contour of the plasma membrane that extended more than a bacterial length, that contained a bacterium at its tip, and in which the plasma membrane apposed the bacterium. The frequency of protrusion formation was expressed as the percentage of intracellular bacteria that were within protrusions. None of the treatments had a significant effect on the number of bacteria present within cells. Results for PtK2 and HeLa cells were similar. In each experiment performed in this study, the confluence of cells was similar under all conditions.
For the comparison of protrusion formation by the ipgB2 mutant with that by the wild-type strain, PtK2 cells were infected with either the wild type or its isogenic nonpolar ipgB2::kan mutant at an MOI of 20, and infection was allowed to proceed as described above. Cells were fixed and permeabilized as described above. Polymerized actin was labeled with Alexa Fluor 568 phalloidin, and DNA was labeled with DAPI. Images of infected cells were acquired as described below and were analyzed for the total number of intracellular bacteria, the number of intracellular bacteria with actin tails, the number of intracellular bacteria in protrusions, and the lengths of actin tails.
For analysis of the role of RhoA per se in Shigella protrusion formation, PtK2 cells were transfected by electroporation with GFP and either dominant negative RhoA(T19N) or the Myc control plasmid at a ratio of 1:10 and were seeded as described above. Sixteen hours posttransfection, cells were infected with wild-type S. flexneri at an MOI of 10 at 37°C. Following an initial invasion period of 1.5 h, cells were washed, and the infection was allowed to continue for an additional 1.5 h in the presence of 50 μg/ml gentamicin. Cells were fixed and permeabilized as described above. Polymerized actin was labeled with Alexa Fluor 568 phalloidin, and DNA was labeled with DAPI. Images were acquired of transfected cells, identified by green fluorescence, that were infected, for three independent experiments. Images were analyzed for the total number of intracellular bacteria and the number of intracellular bacteria in protrusions, defined as described above.
To test whether inhibition of Dia had an effect on bacterial entry into cells, the efficiency of bacterial entry was determined, essentially as described previously (41). PtK2 cells were transfected with either EGFP or DID-EGFP and were seeded as described above. Sixteen hours posttransfection, cells were infected with wild-type S. flexneri at an MOI of 100 at 37°C. Following an initial invasion period of 30 min, cells were washed, and the infection was allowed to continue for an additional 30 min in the presence of 50 μg/ml gentamicin. Cells were fixed as described above, and DNA was labeled with DAPI. Images of transfected cells, identified by green fluorescence, were acquired and analyzed for the presence of intracellular bacteria. At least 30 transfected cells were analyzed for each condition, in each of three independent experiments.
Two distinct assays were used to assess the efficiency of bacterial spread from one cell into adjacent cells: (i) a spreading assay, which assesses spread during the first 3 to 4 h of infection, and (ii) a plaque assay, which assesses spread during the first 48 to 72 h of infection. For analysis of the inhibition of intercellular spread by the DID, a spreading assay was used, because the DID-expressing vector could not be efficiently maintained in the monolayer for the 72 h required to set up and conduct a plaque assay (data not shown). PtK2 cells were transfected by electroporation with DsRed, DID-DsRed, or DID(A256D)-DsRed as described above and were seeded at 70% to 90% confluence. Sixteen hours posttransfection, cells were infected with wild-type S. flexneri carrying pGFPmut2 at an MOI of 0.5 at 37°C; the low MOI was chosen so as to maximize the likelihood that each infectious focus was the result of the initial infection of only a single cell, and not of multiple adjacent cells. Following an initial invasion period of 1 h, cells were washed with fresh DMEM supplemented with 10% fetal bovine serum, 100 μg/ml ampicillin (to maintain pGFPmut2), and 50 μg/ml gentamicin, and the infection was allowed to continue for an additional 2.5 h. Cells were fixed and labeled with DAPI as described above. Transfected cells were identified by red fluorescence. Images were acquired of transfected cells that were highly likely to be the first cell within the field that had been infected, based on (i) the low MOI, (ii) the presence of >5- to 10-fold more bacteria in that cell than in adjacent cells, and (iii) the central location of the cell within the focus of infected cells. Images were analyzed for the number of cells within the focus that were infected. For each experimental condition in each experiment, 6 to 20 infectious foci were analyzed.
For analysis of the intercellular spread of wild-type S. flexneri in HeLa cells depleted or not depleted of Dia1 by RNAi, or for analysis of the intercellular spread of the ipgB2 mutant compared to that of the wild-type strain in untreated HeLa cells, a plaque assay was used. Confluent monolayers of cells grown in 60-mm-diameter dishes or 6-well plates were infected at an MOI of 0.001 at 37°C. For the RNAi-treated monolayers, the cells had been transfected with RNAi using HiPerFect (Qiagen) 48 h prior to infection. No independent RNAi control was used for this set of experiments. Following an initial invasion period of 15 min to 1.5 h, monolayers were washed with fresh MEM and overlaid with 0.5% agarose in DMEM supplemented with 10% fetal bovine serum and 25 μg/ml gentamicin. Forty-eight hours later, monolayers were stained with neutral red, and images of the infected monolayers were acquired using an Epson Perfection 4990 Photo desktop scanner and Adobe Photoshop Elements software (version 2.0). The area of individual bacterial plaques within the monolayers was measured using IPLab software (Scanalytics).
For analysis of the dependence of Dia1 recruitment on the presence of IpgB2, HeLa cells were transfected with Myc or Myc-Dia1 and were seeded as described above. Sixteen hours posttransfection, cells were infected either with wild-type S. flexneri or with the ipgB2 mutant at an MOI of 10, as described above. Following a 3-h infection, cells were fixed, permeabilized, and labeled with DAPI as described above. Labeling of the Myc tag was performed as described above.
Transient depletion of Dia1 and Dia2 was performed in HeLa cells. Dia1 mRNA levels were depleted using either pSUPER-retro-mDia1KD1 and pSUPER-retro-mDia1KD2 shRNA vectors in a 1:1 combination or Dharmacon siGENOME siRNA D-010347-01, while Dia2 RNA levels were reduced using either the V2HS_73202 shRNA vector (OpenBiosystems) or Dharmacon siGENOME siRNA D-018997-02-0002. For experiments in which Dia1 or Dia2 was depleted using shRNA constructs, cells were seeded at a density of 5 × 105 per well in 6-well plates and were allowed to recover overnight in MEM supplemented with 10% fetal bovine serum, nonessential amino acids, and penicillin-streptomycin solution. The following day, 2 μg of the appropriate shRNA vector(s) was added to the cells by using the Arrest-In transfection reagent (OpenBiosystems) according to the manufacturer's protocol. Following a 40-h recovery, transfected cells were selected for 2 days by addition of 1 μg/ml puromycin to the medium. Selected cells were reseeded onto acetone-rinsed glass coverslips and were maintained in MEM supplemented with 10% fetal bovine serum, nonessential amino acids, and 0.2 μg/ml puromycin. For experiments in which Dia1 or Dia2 was depleted using siRNA, 5.7 × 105 cells were transfected in 6-well plates using HiPerFect. Sixteen (Dia2) or 40 (Dia1) hours later, transfected cells were seeded onto glass coverslips and allowed to recover overnight. Cells were infected, fixed, labeled, and imaged as described above, except that an MOI of 10 was used for the infection. Images of infected cells were acquired and analyzed as described above. For each experimental condition in each experiment, 10 or more infected cells were analyzed.
Time lapse microscopic imaging and determination of bacterial speed were performed on semiconfluent monolayers of HeLa cells that had been transfected 16 h earlier with DID-EGFP or EGFP alone, as described above. Transfected cells were infected with wild-type strain 2457T (MOI, 200), and images were recorded at a rate of 1 every 5 s for 5-min periods, as described previously (40). For each experimental condition in each experiment, speeds were determined for 7 or more moving bacteria.
Levels of Dia1 and Dia2 in cells were determined by immunoblotting of whole-cell protein preparations. Adherent cells from 2 wells of a 6-well plate were washed with cold phosphate-buffered saline (PBS) and lifted with 0.05% trypsin in MEM. Cells were recovered by centrifugation, washed twice with cold PBS, and resuspended in 30 μl lysis buffer (50 mM HEPES, 4% sodium dodecyl sulfate [SDS], 300 mM NaCl, 1 mM EDTA, 5 μg/ml aprotinin, 5 μg/ml leupeptin, 1 μg/ml pepstatin A [pH 7.5]). Lysates were boiled for 5 min and incubated for 1 min at room temperature. Then 15 mM N-ethylmaleimide was added, and samples were incubated for 5 min at room temperature. Protein samples were diluted into SDS-polyacrylamide gel electrophoresis loading buffer supplemented with 500 mM NaCl, 2 M urea, and 70 mM β-mercaptoethanol, were separated on 7.5% polyacrylamide gels, and were transferred to nitrocellulose membranes. Separate membranes were probed with antibodies raised against murine Dia11-548 or Dia21-520 and with secondary antibodies conjugated to horseradish peroxidase. β-Actin was detected using a horseradish peroxidase-conjugated anti-β-actin antibody (Sigma). Signal was detected using SuperSignal West Pico chemiluminescent substrate (Thermo Scientific).
PtK2 cells were transfected by electroporation with Myc-Dia1, Myc-DID, Myc-DID(A256D), or Myc alone, as described above. Sixteen hours posttransfection, cells were infected with wild-type S. flexneri or the conditional virB derivative of 2457T at an MOI of 20 to 100, as described above. For the conditional virB strain, 0.1 mM isopropyl-β-d-thiogalactopyranoside (IPTG) was present in the growth medium until the initiation of infection, when it was either removed or maintained at the same concentration for the duration of the infection. After 1.0 to 1.5 h of initial invasion and 1.0 to 1.5 additional hours of infection in the presence of gentamicin, cells were fixed and permeabilized as described above. Polymerized actin was labeled with BODIPY FL phallacidin, and DNA was labeled with DAPI. Labeling of the Myc tag was performed as described above. Results for infection with the wild-type strain were similar for PtK2 and HeLa cells; results for infection with the conditional virB strain were not compared.
Epifluorescence and phase-contrast microscopy were performed using a Nikon Eclipse TE300 microscope equipped with Chroma Technology filters and a Photometrics CoolSNAP HQ charge-coupled device camera (Roper Scientific). Images were acquired using IPLab software. Color figures were assembled by separately capturing images with each of the appropriate filter sets and digitally pseudocoloring the images using Adobe Photoshop software. The statistical significance of differences between experimental results was determined using Student's t test.
In resting cells, the Diaphanous-related formins are present in an autoinhibited state (13), in which the Diaphanous inhibitory domain (DID) binds the Diaphanous autoregulatory domain (DAD) (Fig. (Fig.1A)1A) (26). Isolated Dia1 DID (amino acids 129 to 369) inhibits the actin nucleation activity of Dia1 in vitro (26). We tested whether Dia1 was required for the generation of plasma membrane protrusions by S. flexneri by comparing the percentage of intracellular bacteria within protrusions in cells expressing DID-EGFP to that in cells expressing EGFP alone. We performed these analyses in PtK2 cells, because their flat morphology facilitates the identification of protrusions; similar results were obtained with HeLa cells (data not shown). In the control infections, approximately 15 to 20% of bacteria were found in plasma membrane protrusions at the time the analysis was performed (3 h of infection). Expression of the DID led to a fourfold reduction in the formation of protrusions by S. flexneri (P = 0.002) (Fig. 1B and C; Table Table1)1) but had no significant impact on the frequency or length of actin tails or on the speed of moving bacteria within the cell body (Table (Table1).1). In cells expressing the DID, bacteria accumulated at the cell periphery, with small amounts of polymerized actin at the bacterial pole farthest from the membrane, but were prevented from pushing out against the membrane (Fig. (Fig.1B,1B, arrowheads). Thus, expression of the DID had no significant effect on the movement of bacteria in the cell body or on their accumulation at the cell periphery but inhibited the formation of protrusions by bacteria at the cell periphery.
Not infrequently, in cells expressing the DID, clusters of bacteria were found within cellular projections that resemble retraction fibers (Fig. (Fig.2);2); these projections were distinct from typical bacterial protrusions, both because they contained multiple bacteria and because there were no actin tails associated with these bacteria. Bacteria within protrusions are almost universally associated with actin tails. The presence of retraction fibers in cells in which the RhoA pathway has been blocked is not unprecedented; RhoA activation is thought to be required for retraction of the cell tail during cell migration (7). Moreover, retraction fibers have been observed to trail cells that were induced to migrate by inhibition of RhoA activation of Dia1 by the vaccinia virus protein F11L (46). Thus, the presence of structures resembling retraction fibers in cells expressing the DID construct suggests that the DID is indeed blocking the RhoA pathway by inhibiting the RhoA effectors Dia1 and/or Dia2.
The entry of bacteria was not inhibited by expression of the DID; instead, entry into cells expressing DID was slightly but insignificantly increased over entry into cells expressing GFP alone (61.3% ± 13.0% of cells expressing the DID were infected, versus 40.0% ± 10.0% of cells expressing GFP only; P = 0.2). Therefore, the reduction in the rate of protrusion formation in cells expressing the DID is not due to less-efficient entry of the bacteria into these cells. The expression of a derivative of the DID that is defective in binding to the Dia1 DAD [DID(A256D)] (34) had no effect on bacterial protrusion formation (Fig. (Fig.1D;1D; Table Table1),1), indicating that the inhibitory effect of the DID depends on its interaction with the DAD.
We tested whether a reduction in Dia1 expression via RNAi would similarly inhibit this process. We performed these experiments on HeLa cells with RNAi that targets human Dia. Depletion of Dia1 (Fig. (Fig.3B)3B) led to a significant reduction in protrusion formation (P = 0.02) (Fig. (Fig.44 and Table Table2),2), confirming the role of Dia1 in this process. Since autoinhibition due to the interaction of the DID with the DAD is common to both Dia1 and Dia2, and the Dia1 DID binds the Dia1 DAD and the Dia2 DAD with similar KD (equilibrium dissociation constants) (47), we also tested whether Dia2 could contribute to protrusion formation. Western blot analysis revealed that Dia1 is expressed at approximately equivalent levels in HeLa and Ptk2 cells and that Dia2 is also expressed in HeLa cells (Fig. (Fig.3A).3A). As in the DID inhibition experiments (described above), approximately 15 to 20% of bacteria were found in plasma membrane protrusions of control cells at the time the analysis was performed (3 h of infection). Depletion of Dia2 led to a reduction in protrusion formation that was comparable to that following depletion of Dia1 (P = 0.02) (Fig. (Fig.44 and Table Table2),2), even though the depletion by RNAi was specific for each (Fig. (Fig.3B).3B). Depletion of both Dia1 and Dia2 together led to a slightly smaller reduction in the efficiency of protrusion formation (Table (Table2),2), perhaps because, for reasons that are unclear, the degree of depletion of Dia2 was not as marked as when it was depleted alone (Fig. (Fig.3B).3B). These results did not appear to be an artifact of the specific RNAi construct, since depletion of Dia1 and depletion of Dia2 with independent RNAi constructs that target distinct sites on the mRNA gave similar reductions in protrusion formation: Dia1 depletion led to a 40% reduction (P = 0.03), and Dia2 depletion led to a 47% reduction (P = 0.02), in protrusion formation (Table (Table2).2). Thus, Dia1 and Dia2 are required by intracellular S. flexneri for the efficient formation of plasma membrane protrusions.
The formation of protrusions is thought to constitute a key step in Shigella spread. To test whether the decrease in protrusion formation that occurs upon expression of the DID correlates with a decrease in intercellular spread, we measured the efficiency of movement of S. flexneri from an infected cell to adjacent uninfected PtK2 cells during the first 3.5 h of infection (Fig. 5A to C). We used an assay with which we could selectively analyze the spread from cells that express the transfection construct of interest (45), allowing us to compare the spread from cells expressing the DID to that from control cells. At 3.5 h after the initiation of bacterial infection, the infected monolayers were fixed and stained; transfected cells that were the initial site of bacterial entry were identified; and the percentage of intracellular bacteria that had moved from the site of entry into adjacent cells was quantified (see Materials and Methods). The spread from DID-DsRed-expressing cells (0.5 ± 0.4 infected cell in the infectious focus, excluding the primarily infected cell) was threefold lower than the spread from DsRed-expressing cells (1.5 ± 0.3 infected cells) (P = 0.03), in a manner dependent on DID binding to Dia. The spread from cells expressing DID(A256D)-DsRed (1.3 ± 0.2 infected cells) was significantly different from the spread from DID-DsRed-expressing cells (P = 0.04). (Values are means ± standard deviations [SDs] for 6 to 20 foci of infection per experiment, from three independent experiments). Thus, DID inhibition of Dia reduces both intercellular spread and protrusion formation. The observed correlation between protrusion formation and spread suggests that the former is an important prerequisite for the latter.
We extended these results by showing that depletion of Dia leads to a significant reduction in the spread of S. flexneri through a cell monolayer over 48 h of infection. The area of bacterial spread through monolayers of HeLa cells that had been depleted of Dia1 by RNAi, as determined by a plaque assay, was significantly smaller than that through monolayers of cells treated with a control RNAi (area of spread, 72% ± 5% of the control area; P = 0.009) (Fig. (Fig.5D).5D). These results indicate that efficient intercellular spread of S. flexneri depends on Dia1 and that Dia1 functions in the step of bacterial movement from an infected cell into adjacent cells.
Consistent with a role for Dia1 in protrusion formation by S. flexneri, we found that Dia1 was prominent in actin tails behind bacteria found in protrusions (Fig. 6A to C, arrows). On bacteria within the body of the cytoplasm, the signal from Myc-Dia1 was seen as a thin rim around the bacteria; this signal tended to be more prominent on bacteria that were in the periphery of the cell (Fig. 6A and B, arrowheads). The signal was also seen in a thin rim around protrusions engulfed by adjacent cells (not shown), suggesting that Dia1 is present in the membrane surrounding the bacterium within the engulfed protrusion. Myc-DID also colocalized as a thin rim around S. flexneri at the periphery of the cell (Fig. (Fig.6D,6D, arrowheads), suggesting that Myc-DID localizes to these sites by binding Dia and that it inhibits protrusion formation by blocking the activation of Dia at these sites. Myc-DID(A256D), which is defective in the binding of Dia (34), also localized to the actin tails on bacteria within protrusions but was less prominent than Dia or the DID around bacteria within the body of the cytoplasm (Fig. (Fig.6E),6E), consistent with the observed decrease in the binding of this mutant to Dia. The presence of Myc-DID(A256D) in actin tails within the protrusions may be due to a high concentration of Dia at these sites, combined with the low affinity of DID(A256D) for Dia, or to an interaction with another protein within the protrusions. When cells transfected with a vector expressing Myc alone were infected, the signal from Myc showed no localization to bacteria or to bacterial actin tails (not shown). This pattern of DID localization is consistent with its colocalization to sites of Dia activation by intracellular S. flexneri during protrusion formation and suggests that the DID acts locally at these sites to inhibit Dia-dependent protrusion formation.
Many Shigella proteins that interact with host factors are secreted into the host cytoplasm by the bacterial type III secretion system (12). To investigate a potential role of type III secreted proteins in the recruitment of Dia to intracellular S. flexneri, we infected cells expressing Myc-Dia1 with an S. flexneri derivative that conditionally expresses the type III secretion machinery. In this strain, the global regulator of type III secretion, VirB, is expressed under the control of an IPTG-inducible promoter (38). Since the entry of Shigella into cells depends on type III secretion, IPTG was included in the medium until the time of infection and then was either removed or maintained for the duration of the infection. Under these conditions, VirB expression becomes undetectable within 25 min of washout of the inducer (25). At 2 h of infection, the recruitment of Myc-Dia1 to intracellular S. flexneri was indistinguishable under non-VirB-expressing and VirB-expressing conditions (data not shown), suggesting that Dia recruitment either is independent of type III secretion or is dependent on a bacterial effector that is secreted early during infection.
In the stress fiber formation pathway, Dia functions as a downstream effector of activated RhoA (18, 32, 46). Given that Dia activity is required for Shigella protrusion formation, it was possible that Shigella activation of Dia during protrusion formation might depend on RhoA. To test this, we compared the efficiency of protrusion formation in PtK2 cells expressing a Myc-tagged dominant negative form of RhoA [RhoA(T19N)] to that in cells expressing Myc alone. The expression of RhoA had no effect on protrusion formation [17.3% ± 2.8% of bacteria were in protrusions in RhoA(T19N)-transfected cells versus 14.7% ± 2.0% of bacteria in Myc-transfected cells (P = 0.5)]. The dominant negative RhoA construct was highly expressed, as determined by Western blot analysis, and the expression of this construct had a dominant negative effect on stress fiber formation: cells transfected with the RhoA(T19N) construct displayed substantially fewer stress fibers than cells transfected in parallel with either a wild-type or a constitutively active RhoA construct (data not shown). These results suggest that activation of RhoA is not required for Shigella protrusion formation.
Shigella proteins that are translocated into cells via the type III secretion system display diverse effects on host cell processes that enhance pathogenesis. Shigella IpgB2, a protein secreted by the type III secretion system, has been shown to bind cellular Dia1 in coimmunoprecipitation assays (3). In addition, expression of IpgB2 in mammalian cells induces the formation of stress fibers (Fig. 7A and B) (3). Together these findings suggested that IpgB2 may modulate Dia1 to enhance S. flexneri protrusion formation during intercellular spread. However, the role of IpgB2 in protrusion formation by, and spread of, Shigella spp. was not tested in this earlier study.
We tested whether S. flexneri lacking IpgB2 would be defective in protrusion formation or intercellular spread. Bacterial spread, as assessed by the formation of plaques in HeLa cell monolayers during 48 h of infection, was similar for the wild-type strain and an isogenic nonpolar ipgB2 deletion mutant (the diameter of spread, given as the mean ± SD, was 0.95 ± 0.17 mm for the wild type and 0.87 ± 0.20 mm for the ipgB2 mutant [Fig. [Fig.7]).7]). In addition, the frequencies of protrusion formation (means ± SDs, 39% ± 8% for the wild type and 32% ± 15% for the ipgB2 mutant) and the lengths of the protrusions (means ± SDs, 5.0 ± 1.0 μm for the wild type and 4.6 ± 0.9 μm for the ipgB2 mutant) were similar for the two strains. Moreover, overexpression of IpgB2 by transfection of cells expressing the DID did not rescue the efficiency of protrusion formation (data not shown).
We examined whether the recruitment of Dia1 to intracellular bacteria was dependent on IpgB2. We found that Dia1 colocalized with an ipgB2 mutant in a manner similar to its colocalization with wild-type S. flexneri (Fig. 7D and E). In both cases, the signal from Myc-Dia1 was prominent around bacteria at the periphery of the cell and in actin tails within bacterial plasma membrane protrusions. The signal from HeLa cells expressing Myc alone did not localize around intracellular bacteria (not shown), indicating that the signal observed around the bacteria in cells expressing Myc-Dia1 was specific. Thus, IpgB2 is not required for the recruitment of Dia1 to intracellular bacteria.
Therefore, while IpgB2 is sufficient to induce stress fiber formation, likely as a result of its previously described interaction with Dia1 (3), it is not essential for Dia recruitment, Dia1-dependent formation of protrusions, or intercellular spread by S. flexneri. Although IpgB2 is not required for enhancing intercellular spread, it is possible that one or more other Shigella proteins may be functionally redundant with IpgB2 in this pathway. Although we have no evidence that a redundant protein exists, functional redundancy is common in bacterial type III secretion systems (50).
The spread of intracellular Shigella from an infected cell into an adjacent cell has long been thought to depend on the formation of actin-based bacterial protrusions from the surface of the infected cell. Our findings constitute the first demonstration that activation of Dia is required for the formation of actin-based cell surface projections and for spread during microbial infection. Moreover, the observation that intercellular spread is significantly diminished when protrusion formation is inhibited establishes a direct correlation between protrusion formation and intercellular spread.
The role of Dia1 in enhancing Shigella protrusion formation may reflect its function in the maintenance and remodeling of the cellular cortical actin network. The cell cortex, lying just beneath the plasma membrane, contains a dynamic and dense network of actin filaments. The cortical actin network is continually remodeled, and its remodeling is critical to the maintenance of cell shape and to cell motility. Actin polymerization in the cell cortex is regulated by RhoA through its effectors Dia1 and Dia2 (1, 39, 48). Our results may reflect a role of Dia in enhancing Shigella protrusion formation through increased remodeling of the cortical actin network. Based on our findings, we suggest a model in which Shigella directs the reorganization of cortical actin into an orientation that is perpendicular to the plasma membrane and utilizes the force generated by the formation of parallel arrays of polymerized actin to push out against the plasma membrane.
In addition to promoting remodeling of the actin cortex, the role of Dia in protrusion formation may also be due to a direct role in force generation at the plasma membrane. Formins generate 1.3 pN or more of force per actin filament (22). Forces of tens of pN are thought to be required to enable a filopodium to protrude against the resistance of the plasma membrane (9). Such a force could be generated by formins associated with a bundle of actin filaments. To generate a plasma membrane protrusion, bacteria must gather similar force using the resources of the cell. Our data are consistent with a model in which the formin Dia1 is responsible for generating part or all of the force that is required in this process during S. flexneri infection.
Shigella-induced actin polymerization in the cell body depends on localized activation of the Arp2/3 complex by N-WASP at the bacterial surface (11, 14). Our results demonstrate that whereas Dia is not required for actin tail assembly in the cell body, it is required for the efficient formation of protrusions and for intercellular spread. Moreover, Dia1 recruitment to the bacteria is enriched at the cell periphery (Fig. (Fig.66 and Fig. 7D and E). These findings indicate that when the bacteria reach the cell cortex or plasma membrane, a switch likely occurs, leading to the activation of Dia-mediated actin polymerization. This is similar to a switch that occurs during vaccinia virus infection. Vaccinia virus moves from the peri-Golgi region to the cell periphery by microtubule-based motility (33); at the plasma membrane, viral particles switch from microtubule-based motility to actin-based motility (29). How Shigella induces this switch and which bacterial proteins are involved in this process will be the subject of further investigation.
The process of Shigella spread from one cell into an adjacent cell involves the uptake of a plasma membrane-bound bacterial protrusion by the adjacent cell in a process that resembles macropinocytosis (37). It is possible that, in addition to playing a role in Shigella protrusion formation, Dia is involved in the uptake of the protrusion by the adjacent cell. Our results do not directly address this. Moreover, although it is not known whether the uptake by the adjacent cell mechanistically mimics the initial entry of the bacterium into cells from the extracellular milieu, our data on entry indicate that Dia has little or no role in initial entry.
In sharp contrast to the requirement for activation of Dia in S. flexneri spread is the requirement for inhibition of the RhoA-Dia pathway for productive infection with vaccinia virus. The vaccinia virus protein F11 inhibits the RhoA-Dia pathway and enhances the release of vaccinia virus from cells (4, 46). Moreover, expression of constitutively active RhoA or Dia inhibits viral particle accumulation at the cell periphery, actin tail formation on viral particles at the cell periphery, and viral release from cells (4).
The observation that divergent effects on the RhoA-Dia pathway can lead to similar outcomes, namely, the exit of microorganisms from the cell, suggests that the two organisms have evolved distinct mechanisms for manipulating the cortical cytoskeleton during spread. Specific differences in how the microorganisms move to the cell periphery may be at the core of the differences in these mechanisms of exit. Vaccinia virus moves to the cell periphery by microtubule-based motility (33), whereas Shigella moves to the periphery by polymerization of actin tails (5, 24). The findings of Arakawa et al. (4) suggest that vaccinia virus inhibition of the RhoA-Dia pathway induces a reorganization of the cortical cytoskeleton that facilitates the movement of the virus through the cortical cytoskeleton to the plasma membrane. Our data suggest a model in which, once Shigella arrives at the plasma membrane via N-WASP-dependent actin-based motility, its activation of the RhoA-Dia pathway enables Dia-dependent actin polymerization and reorganization of the actin cortex in such a way as to enable the bacteria to push outward from the cell surface and into adjacent cells.
We thank D. Colon for technical assistance and B. Cormack, H. N. Higgs, R. R. Isberg, L. Tsiokas, and N. Watanabe for providing reagents.
This research was supported by National Institutes of Allergy and Infectious Diseases grants AI052354 (to S. B. Snapper and M.B.G.), AI073967 (to M.B.G.), and AI081724 (to M.B.G.), by a Harvey Fellowship from the Mustard Seed Foundation (to J.E.H.), and by funds from the Executive Committee on Research of the Massachusetts General Hospital (to M.B.G.).
Editor: J. B. Bliska
Published ahead of print on 19 October 2009.