ACD toxin-produced actin oligomers poison formin-controlled actin polymerization

Abstract

The actin crosslinking domain (ACD) is an actin-specific toxin produced by several pathogens, including life-threatening spp. of Vibrio cholerae, Vibrio vulnificus, and Aeromonas hydrophila. Actin crosslinking by ACD is thought to lead to slow cytoskeleton failure owing to a gradual sequestration of actin in the form of nonfunctional oligomers. Here we found that ACD converted cytoplasmic actin into highly toxic oligomers that potently “poisoned” the ability of major actin assembly proteins, formins, to sustain actin polymerization. Thus, ACD can target the most abundant cellular protein by employing actin oligomers as secondary toxins to efficiently subvert cellular functions of actin while functioning at very low doses.

Bacterial toxins are the deadliest compounds on the planet. As little as a single molecule of a delivered toxin can compromise vital functions or even kill an affected host cell (1, 2). This is achieved by amplification of a toxin enzymatic activity via signaling cascades (e.g. by cholera, pertussis, and anthrax toxins) or via enzymatic inhibition of vital host complexes present in relatively few copies (e.g. Shiga and diphtheria toxins acting on ribosomes). Such efficiency is crucial because i) the amount of a toxin produced early upon infection is limited by an initially small number of bacterial cells; ii) the host is protected by commensal bacteria; and iii) the host immune system efficiently neutralizes toxins by means of adaptive (antibodies) and innate (e.g. defensins) (3) humoral defense factors.

Owing to its importance in multiple cellular processes, actin is a common target for bacterium- and parasite-produced toxins. Upon delivery to the cytoplasm of host cells via Type I (as part of MARTX toxin) (4) or Type VI (within VgrG1 toxin) (5) secretion systems, the actin crosslinking domain toxin (ACD) catalyzes the covalent crosslinking of K50 in subdomain 2 of one actin monomer with E270 in subdomain 3 of another actin monomer via an amide bond, resulting in the formation of actin oligomers (6, 7). The actin subunits in the oligomers are oriented similar to short-pitch subunits in the filament, except that a major twist of the subdomain-2, required to accommodate such orientation, disrupts the normal inter-subunit interface and precludes polymerization (6).

The currently accepted mechanism of ACD toxicity, via sequestering of bulk amounts of actin as non-functional oligomers, is compromised owing to the high concentration (hundreds of micromolar) of actin in a typical animal cell. Extrapolation of in vitro determined rates of the ACD activity (7) to cellular conditions suggests that a single ACD molecule per cell (i.e. ~ 1 pM) would require over six months to covalently crosslink half of all cytoplasmic actin.

In contrast to these estimations, the integrity of the intestinal cell monolayers was disrupted when only a small fraction of cellular actin (2-6%) was crosslinked by ACD (Fig. 1A-C; fig. S1). To account for such dramatic effects, we hypothesized that the ACD-crosslinked actin oligomers are highly toxic because they can exert an abnormally high affinity to actin-regulatory proteins containing several actin-binding domains. To identify potential high-affinity partners of the actin oligomers, anthrax toxin delivery machinery was used to deliver ACD (8) into HeLa cells transfected with double-tagged (Twin-Strep-tagII and hemagglutinin) actin (SHA-actin; fig. S2) and used for a pull-down assay. Several formins (DIAPH1, DIAPH2, DAAM1, and INF2) preferentially bound to the ACD-crosslinked actin oligomers (Fig. 1D). Treatment of epithelial monolayers with the formin inhibitor SMIFH2 affected the monolayer integrity similar to ACD, whereas the Arp2/3 complex inhibitor CK-666 did not (fig. S3).

Figure 1. Integrity of intestinal monolayers is compromised by low concentration of actin oligomers.

(A-C) Transepithelial electrical resistance (TEER) of IEC-18 monolayers (A) was assessed upon cytoplasmic delivery of LF_NACD or a catalytically inactive mutant as a control and correlated with the accumulation of ACD-crosslinked actin species by anti-actin immunoblotting (B) and cell morphology (C). Additional antiactin blots and quantitation of crosslinked actin are presented on fig. S1. (D) SHA-actin pull-down. Lanes A: SHA-actin transfected cells treated with inactive LF_NACD (non-crosslinked actin). Lanes X: SHA-actin transfected cells treated with active LF_NACD (crosslinked actin). Lanes C: non-transfected untreated cells used as a negative control. “NaCl” and “FA” – fractions eluted from Strep-Tactin beads with 0.5 M NaCl and 50% formamide, respectively. Samples were subjected to immunoblotting and probed with anti-HA, anti-actin, various anti-formin, and anti-profilin antibodies.

Formins are a major family of actin assembly factors involved in numerous actindependent cellular processes. The major functional domains of formins, formin homology domains 1 (FH1) and 2 (FH2), cooperate in nucleation and elongation of actin filaments. A non-covalent FH2/FH2 homodimer nucleates and remains at the polymerizing barbed end to facilitate processive filament elongation while protecting the filament from capping (9). Tandem poly-proline stretches within the FH1 domains bind profilin-actin complexes and accelerate elongation by as much as 10-fold (10-12). FH1 domains of all formins preferentially bound to the oligomers (Fig. 1D) contain 4-14 tandem poly-proline (PP) stretches, which may contribute to strong profilin-mediated interaction with the oligomers.

To elucidate the mechanism of formin inhibition, we employed constitutively active FH1-FH2 fragments of mDia1 and mDia2 (mouse orthologues of human DIAPH1 and DIAPH3, respectively) to monitor actin polymerization at the individual filament level by total internal reflection fluorescence microscopy (TIRFM; Fig. 2, 3; fig. S4). In the presence of human profilin-1 (PFN1), the oligomers caused very prominent reversible blocks of elongation of formin-controlled, but not formin-free actin filaments (Fig. 2A-F; fig. S4B,C; Movies S1-5). Formin-controlled filaments were identified by faster growth with a dimmer appearance (Fig. 2A,E) (10), or via direct labeling of formin (Fig. 3A).

Figure 2. Effects of ACD-crosslinked actin oligomers on polymerization of individual filaments controlled by mDia1(14PP).

(A) mDia1(14PP)-mediated polymerization from profilin-actin complexes in the absence (top) and presence (bottom) of actin oligomers (A-Oligo) was monitored by TIRFM. Arrowheads denote actin barbed ends: green – mDia1-controlled (dim and fast); yellow – mDia1-free (bright and slow); red – mDia1-controlled stopped by the oligomers. (B, C) Quantitation of (A): filament elongation plots in the presence (B) or absence (C) of PFN1. Green and red curves describe filament elongation in the absence and presence of oligomers, respectively. Arrows denote the beginning and arrowheads indicate the end of elongation blocks caused by the oligomers on representative curves highlighted in black. (D) IC₅₀ of oligomers determined by TIRFM as percent of stopped filaments (black) or growth rate inhibition (red curves). (E) TMR-labeled actin (red) was polymerized in the presence of mDia1(14PP) and PFN1 without oligomers followed by flow of Oregon Green (OG) actin, oligomers, and PFN1. Arrowheads are as for (A). (F) Quantitation of (E): growth of mDia1-controlled filaments (green traces) and mDia1-free filaments (yellow traces). Better polymerization properties of OG-actin result in faster elongation at the formin-free ends.

Figure 3. Effects of ACD-crosslinked actin oligomers on polymerization of individual filaments controlled by mDia2 and mDia1 with various FH1 lengths.

(A) OG-actin (green) polymerization in the presence of SNAP-549-mDia2 (red) and PFN1 before and after the addition of oligomers (black arrow) was monitored by TIRFM. Red arrowheads indicate SNAP-549-mDia2 at an actin filament; white arrowheads indicate formin-free filament. Kymograph shows a stalled SNAP-549-mDia2-controlled filament upon addition of oligomers. (B) Effects of oligomers on formin-free filament elongation and elongation controlled by mDia2 and mDia1 formins with various FH1 lengths: 14PP, 2PP, and FH2 (no PP-stretches). (C, D) Oligomer association (k_on) (C) and dissociation (k_off) (D) rates for mDia1(14PP) and mDia2.

In the presence of PFN1, the fraction of blocked mDia1 formin-associated filaments as well as the inhibition of averaged growth rates depended on the concentration of the added oligomers with an IC₅₀ of 1.2 ± 0.6 (SEM) nM (Fig. 2D), in good agreement with the apparent equilibrium inhibition constant determined kinetically (_appK_i = k_off/k_on = 2.5 nM; Fig. 3C,D). After stops (oligomer dissociation), the filaments continued to polymerize with the rates characteristic for formin-controlled filaments (Fig. 2B; fig. S4A). In the absence of PFN1, the inhibition appeared to occur via a similar mechanism, but the overall effect was weaker and the average duration of the blockage events was substantially shorter (Fig. 2C,D). Although a profilin-mediated interaction of the oligomers with the PP stretches of FH1 was not absolutely required, it strongly amplified the efficiency of the inhibition at the elongation stage by contributing to multisite interaction with the oligomers. Thus, mDia1 constructs (fig. S5A) with either removed FH1 domains (FH2 only) or shortened from fourteen (14PP) to two PP-stretches (2PP) showed progressively lower response to inhibition by the oligomers in the presence of PFN1 (Fig. 3B). Similarly, the _appK_i of oligomers for mDia2 (containing two PP-stretches) was 7.5 fold higher than that for mDia1 and depended on PFN1 (Fig. 3B-D).

The inhibition of formin-mediated polymerization measured at the individual filament level correlated well with the inhibition observed in bulk pyrene assays (Fig. 4; fig. S5,S6). During spontaneous polymerization in the absence of PFN1, high concentrations (75-500 nM) of the oligomers mildly accelerated the polymerization, while mild inhibition was observed in the presence of profilin (Fig. 4A,B). This is likely because of a low level incorporation of the oligomers into the filaments (6) in the absence, but not in the presence of PFN1 (fig. S5D), leading to filament severing similar to that observed for actin species with impaired inter-subunit surfaces (13).

Figure 4. Actin oligomers inhibit mDia1-controlled actin polymerization in bulk pyrenyl-actin assays.

(A-D) Effects of actin oligomers (A-Oligo) on actin polymerization in the absence (A, B) or presence of mDia1(14PP) (C, D); without (A, C) or with PFN1 (B, D). Normalized FL – pyrene fluorescence expressed in percent of maximum. (E, F) Inhibition of profilin-dependent and independent actin polymerization controlled by various length FH1 mDia1 constructs (14PP, 5PP, 2PP, or FH2 only; see fig. S5A,B and S6) assessed in the absence (E) and presence of PFN1 (F). (G) Apparent K_i for inhibition of mDia1(14PP) by the oligomers in the presence of PFN1 was calculated by measuring IC₅₀ at two different concentrations of actin.

In contrast, the oligomers potently inhibited actin polymerization directed by mDia1 in the presence and, to a lesser extent, absence of PFN1 (Fig. 4C-F; fig. S6). Fitting the inhibition of actin polymerization at 50% of maximum to a binding isotherm equation resulted in an IC₅₀ for the mDia1(14PP) construct equal to 2.0 ± 0.2 (SEM) nM and 4.8 ± 0.6 (SEM) nM in the presence and absence of PFN1 (Fig. 4E,F). The ACD-crosslinked actin dimers purified to homogeneity (fig. S5B) inhibited the mDia1-controlled polymerization less efficiently than the mixture of higher order oligomers (fig. S5F-H), suggesting that the inhibition is amplified via multivalent interactions of the oligomers with mDia1. Accordingly, shortening the FH1 domain progressively decreased the efficiency of inhibition with the IC₅₀ values reaching ~30 and ~16 nM for the mDia1(FH2) constructs in the presence and absence of PFN1, (Fig. 4E,F; fig. S6).

Kinetic modeling (fig. S8) revealed that inhibition of both nucleation and elongation is required to accurately describe the effects of the oligomers on formincontrolled actin polymerization. Using experimentally determined parameter values for inhibition of elongation, good fits to the data (Fig. 4) could be found by assuming that oligomers also inhibit nucleation by binding to free mDia1(14PP) formin with dissociation constants of 0.8 and 5 nM in the presence and absence of PFN1 (fig. S8D,E). Inhibition of nucleation by the oligomers in the absence of PFN1 was also observed experimentally in filament seeding assays (fig. S7) and TIRFM experiments (fig. S4D-G). Similar experiments in the presence of PFN1 were less conclusive owing to the overall lower nucleation ability of formins under these conditions (fig. S7G,H and fig. S4F,G). To improve accuracy, modeling had to account for filament severing owing to incorporation of the oligomers in the absence of PFN1 (Fig. 4A,C; fig. S8C,D).

Bacterial toxins are well known to disorganize the actin cytoskeleton acting via Rho family GTPase controlled signaling pathways (14). Here we found that toxins can not only exploit existing signaling pathways, but also initiate a new toxicity cascade with de novo produced crosslinked actin species as “second messengers”. Owing a unique combination of properties that is neither present in G-nor F-actin (fig. S9A), these new actin species bind with high affinity to formins and adversely affect both nucleation and elongation abilities of these proteins causing their potent inhibition in profilin-dependent and independent manners (fig. S9B). Thus, ACD creates toxic derivatives of actin with a disruptive “gain of function” mode of operation. We propose that the seemingly straightforward original assumption that ACD acts via the accumulation of bulk amounts of non-functional actin is inaccurate, or at least incomplete. The toxin can be highly efficient at very low concentrations by acting on formins and potentially other actin regulatory proteins. This finding calls for the careful re-evaluation of mechanisms employed by other actin-related toxins, both of protein and small-molecule nature.