Prepore Stability Controls Productive Folding of the BAM-independent Multimeric Outer Membrane Secretin PulD*

Members of a group of multimeric secretion pores that assemble independently of any known membrane-embedded insertase in Gram-negative bacteria fold into a prepore before membrane-insertion occurs. The mechanisms and the energetics that drive the folding of these proteins are poorly understood. Here, equilibrium unfolding and hydrogen/deuterium exchange monitored by mass spectrometry indicated that a loss of 4–5 kJ/mol/protomer in the N3 domain that is peripheral to the membrane-spanning C domain in the dodecameric secretin PulD, the founding member of this class, prevents pore formation by destabilizing the prepore into a poorly structured dodecamer as visualized by electron microscopy. Formation of native PulD-multimers by mixing protomers that differ in N3 domain stability, suggested that the N3 domain forms a thermodynamic seal onto the prepore. This highlights the role of modest free energy changes in the folding of pre-integration forms of a hyperstable outer membrane complex and reveals a key driving force for assembly independently of the β-barrel assembly machinery.

Outer membrane proteins (OMPs) 4 allow Gram-negative bacteria to exchange nutrients and macromolecules with their environment. Classically, periplasmic broad-specificity chaperones deliver OMPs to the ␤-barrel assembly machinery (BAM) that catalyzes OMP OM insertion (1). In contrast, a group of large multimeric OMPs in which all protomers contribute part of their sequence to a single, shared transmembrane pore rely on the lipoprotein outer membrane localization (Lol) system for membrane targeting and on lipid-assisted self-insertion (2). Many OMPs that use this alternative assembly route are secretion portals for virulence factors or for the surface presentation of pili, and thus are potential targets for chemicals that abolish bacterial virulence.
The first OMP shown to assemble independently of BAM was the secretin PulD (3). PulD dodecamerizes to form the OM pore of the Klebsiella oxytoca type II secretion system for the secretion of pullulanase (PulA) (4). Mature full-length PulD (PulD fl ), comprising amino acids 28 to 660, forms a modular structure ( Fig. 1A) with periplasmic flexible arms (N 0 to N 2 ) (5), a core structure that is referred to as an inverted cup-and-saucer (N 3 in the periplasm and the partly membrane-embedded C (Fig. 1B)) (6 -8), and a domain S for binding to the fatty acylated pilotin PulS (9,10), which delivers PulD to the OM (11) via the Lol system (12). The PulD core structure is hyperstable and resists denaturation by SDS, urea, and trypsin. The acquisition of these characteristics over time showed that a PulD variant lacking the flexible arms, PulD 28 -42/259 -660 (where the superscript denotes the numbers of the included amino acids (Fig. 1A)), forms at least one multimeric intermediate (a prepore) prior to membrane insertion (13). The prepore was isolated by a Thr 470 to Ile substitution in the C domain of both PulD 28 -42/259 -660 and PulD fl (14). Prepores are only SDS resistant and have a well structured cup but a collapsed outer chamber (14). Genetic dissection demonstrated that PulD cannot multimerize without N 3 (15) and PulD 28 -42/259 -660 I322S, a PulD 28 -42/259 -660 variant carrying an I322S substitution on the N 3 surface (Fig. 1B), is multimerization defective (16). These results suggest an important role for N 3 as a regulator for membrane insertion. To test this, the role of N 3 in PulD folding was investigated here by domain exchange of N 3 with the homologous N 1 and N 2 domains, by the effect of the I322S substitution on the structural, energetic, and dynamic properties of N 3 and by the formation of mixed multimers between unaltered PulD protomers and PulD protomers containing modifications that affect at least one folding step. The results provide the first insights into the folding energy landscape of an OMP of such large dimensions.
The Effect of the I322S Substitution Is Specific for the N-subdomain Adjacent to the C Domain-Introducing the I322S substitution into PulD fl resulted in a similar effect as observed for PulD 28 -42/259 -660 I322S: PulD fl I322S predominantly migrated as a monomer, but the amount of multimers increased when it was solubilized from the liposomes with ZW 3-14 prior to SDS-PAGE ( Fig. 2B and Table 1). Although some SDS-resistant PulD fl I322S multimer was formed in vitro and in vivo, PulD fl I322S did not support efficient PulA secretion in the presence of all the Pul components in vivo (Fig. 3A). To validate that the secretion defect was not because of inefficient OM targeting of PulD fl I322S, the phage shock protein (Psp) response was measured by the amount of PspA produced upon PulD fl I322S production in the presence or absence of PulS. Typically, PspA levels are high when PulD fl is suggested to insert into the inner membrane in the absence of PulS, whereas they are low when PulS efficiently delivers PulD fl to the OM (11). In the presence of PulS, PulD fl I322S elicited a smaller Psp response than PulD fl WT, consistent with the smaller amount of PulD fl I322S multimers produced when compared with the amount of PulD fl WT multimers (Fig. 3B). Importantly, the amount of PspA produced in the presence of PulD fl WT or PulD fl I322S (but not PulS) was more than halved when either of the PulD fl variants was produced in the presence of PulS, suggesting that both PulD fl variants were targeted to the OM with equal efficiency (Fig. 3, A and B). Hence, the I322S substitution created a folding defect in vitro and in vivo.
Although homologous, replacing N 3 with N 2 and N 1 did not result in efficient PulD folding. A striking difference between the three domains is the length of the L 2 -loop, which is shorter in N 2 than in N 3 and is only a few amino acids in N 1 (Fig. 2D). Deletion of L 2 in N 3 still allowed PulD 28 -42/259 -660 ⌬L 2 to form SDS-resistant multimers, excluding a role for L 2 in folding ( Fig. 2A and Table 1). Surprisingly, introducing I322S in PulD 28 -42/259 -660 ⌬L 2 yielded SDS-resistant PulD 28 -42/259 -660 ⌬L 2 I322S multimers ( Fig. 2A and Table 1).
The I322S Substitution Destabilizes N 3 -To determine the effect of the I322S substitution on N 3 structure and stability, N 3 domains were purified that carried none, one, or both of the I322S and ⌬L 2 modifications. Circular dichroism (CD) spectroscopy indicated that the secondary structure of N 3 was unaffected by any of the modifications made (Fig. 4A). N 3 domains resisted complete unfolding upon thermal denaturation (Fig. 4, A and B). N 3 I322S was the least thermostable with thermostability increasing in the order of N 3 I322S Ͻ N 3 ⌬L 2 I322S Ͻ N 3 WT Ͻ N 3 ⌬L 2 (Fig. 4B).
The I322S Substitution Increases N 3 Dynamics-Retention of the structure in the N 3 variants was confirmed by the exchange The secondary structure elements correspond to those found in the model of N 3 . The N domains were aligned using MAFFT version 7 with the highest gap penalty. The position of Ile 322 is highlighted by the asterisk and the deleted L 2 -loop by the rectangle. The sequence similarity of N 3 with N 1 and N 2 is 36 and 42%, respectively, and sequence identity is 16 and 22%, respectively.

TABLE 1 Quantification of PulD multimer formed upon in vitro synthesis in the presence of lecithin liposomes
The multimer fraction was determined by densitometry of the multimer and monomer bands on immunoblots after SDS-PAGE analysis as shown in Fig. 2

TABLE 2 Quantification of mixed PulD multimers formed upon in vitro synthesis in the presence of lecithin liposomes
The multimer fraction was determined by densitometry of the multimer and monomer bands on immunoblots after SDS-PAGE analysis as shown in Fig. 6. Errors represent standard deviations over 3 independent measurements.

SDS resistant Urea resistant
PulD fl PulD 28 (19,20). Except for the expected loss of 13 fast-exchangeable amino acids in L 2 , neither the L 2 deletion nor the I322S substitution affected the global hydrogen/deuterium exchange (H/DX) behavior of intact N 3 at early time points (supplemental Fig. S3A and Table S1). However, the number of slow exchangeable amide hydrogens was reduced by ϳ30% in N 3 and in N 3 ⌬L 2 by the I322S substitution, whereas the number of intermediate exchangeable ones increased (supplemental Table S1), indicating a change in protein dynamics.
To locate which N 3 segments were affected by the I322S substitution, H/DX in N 3 domains was analyzed by MS after online pepsin digestion (Fig. 5A, supplemental Fig. S4A). Three N 3 I322S segments showed increased deuterium uptake compared with those of N 3 WT. The largest average uptake difference (17%) was observed in a segment represented by peptide Glu 334 -Leu 340 , which forms the C terminus of ␣-helix 2 ( Fig.  5B) and is the most proximal structural element to the C domain in the PulD sequence (Figs. 1 and 2D). The two other segments (e.g. peptides Lys 270 -Leu 282 and Asp 307 -Asp 332 ) had an uptake difference of 7% (Fig. 5A) and form a patch on the N 3 structure opposite to the C terminus of ␣-helix 2 (Fig. 5B). Introducing the I322S substitution in N 3 ⌬L 2 affected the same region of the N 3 domain, but uptake differences were smaller or disappeared (for peptide Ile 322 -Leu 333 ) ( Fig. 5, C-E, and supplemental Fig. S4B). Glu 334 -Leu 340 remained the peptide with the largest uptake difference (9%), but kinetics closely resembled that of N 3 WT (Fig. 5E, far right panel).
H/DX coupled with NMR or MS can correlate local H/DX rates to the global ⌬G 0 of a protein or the global ⌬⌬G 0 between a protein and its variants (21,22). The peptides in Fig. 5E represent each of the segments that showed differential H/DX behavior upon the introduction of the I322S substitution. Excluding Ile 322 -Leu 333 , which is not directly comparable as it contains the site of the substitution, all peptides fitted best to a double exponential ( Fig. 5E and supplemental Table S1). The slowest exchange rates were found in Glu 334 -Leu 340 , which also had the largest uptake difference, suggesting that the C terminus of ␣-helix 2 plays a role in maintaining N 3 stability. ⌬⌬G 0 local were calculated from the three slowest exchangeable hydrogens in the peptides. Hence, for all N 3 domains the slow rates of Glu 334 -Leu 340 were used, whereas the slow rates from the three peptides were used for N 3 I322S as each peptide had one slowly exchanging hydrogen (supplemental Table S1). ⌬⌬G 0 local correlated well with ⌬⌬G 0 global obtained by equilibrium unfolding with a slope of 0.9 (Fig. 4D).  Folding Defects Are Overcome in Mixed PulD Multimers-How does N 3 energetics affect PulD multimer folding? Because multiple domains might need to interact to fold the PulD native multimer, it was assessed whether PulD 28 -42/259 -660 I322S can form mixed multimers with wild-type PulD fl (PulD fl WT) during co-synthesis. As shown before (13), co-synthesis of PulD fl WT and PulD 28 -42/259 -660 produces mixed multimers containing from 0 up to 12 protomers from one form and, reciprocally, from 12 down to 0 of the other form. The mixed multimers form a regular ladder between 540 (a PulD 28 -42/259 -660 dodecamer) and 825 kDa (a PulD fl dodecamer) upon SDS-PAGE (Fig. 6A). The biochemical determinants of the native PulD multimer can be used to assign structural states to the mixed multimers: an SDS-and urea-resistant mixed multimer forms a membrane-inserted PulD 28 -42/259 -660 pore (13), whereas an SDS-resistant but urea-sensitive mixed multimer forms a prepore (as typified by PulD 28 -42/259 -660 T470I (14)). Trypsin resistance of the PulD pore is not useful in this case, because it trims all protomers to the same molecular mass. Ladders produced upon co-synthesis of PulD fl WT and PulD 28 -42/259 -660 were SDS-and urea-resistant, indicative of native PulD structures in the mixed multimers ( Fig. 6A and Table 2). In the presence of PulD 28 -42/259 -660 I322S, SDS and urea resistance required six or more PulD fl WT protomers in the multimer (Fig. 6, A and B, and Table 2). Thus, native structures can form in the presence of the I322S substitution and suggest that either neighboring N 3 WT domains correct the conformation of N 3 I322S or that the N 3 -ring is required to reach a critical stability. The distribution of protomers in a multimer is not easily determined (the N 0 -N 2 domains in PulD fl are not visible by EM). Hence, to distinguish between these possibilities, mixed multimers were produced between PulD 28 -42/259 -660 I322S and PulD fl variants that differ in N 3 stability. PulD 28 -42/259 -660 I322S formed mixed multimers with all PulD fl variants (Fig. 6A). Importantly, SDS-and urearesistant mixed multimers containing PulD fl ⌬L 2 (which carries the most stable N 3 domain) could incorporate up to four extra PulD 28 -42/259 -660 I322S protomers when compared with multimers containing PulD fl WT or PulD fl ⌬L 2 I322S (Fig. 6A and Table 2). Consistent with the poor multimerization of PulD 28 -42/259 -660 I322S and PulD fl I322S, SDS-and urea-resistant mixed PulD 28 -42/259 -660 I322S/PulD fl I322S-multimers were low in abundance ( Fig. 6A and Table 2). Hence, the results mirrored the stability and dynamics of the N 3 domains and support the idea that the stability in the N 3 -ring needs to attain a critical value to enable PulD membrane insertion.
Effect of the I322S Substitution on the PulD Folding Mechanism-To fold into the SDS-resistant prepore, the SDSsensitive PulD 28 -42/259 -660 multimer must overcome a transition barrier, defined by the Arrhenius activation energy (E a ).
Failure to produce an SDS-resistant prepore in PulD 28 -42/259 -660 I322S either results from an insurmountable E a (which relates to the reaction rate constant k via k ϭ Aexp(ϪE a /RT), where A is a constant that depends on the collision frequency) or from destabilization of the SDS-resistant prepore to such an extent that the SDS-sensitive multimer is the more stable structure (supplemental Fig. S5, A-C). Rate constants for SDS-resistant PulD multimerization are measured by the intensities of the multimer bands over time upon SDS-PAGE (13). Although these are easily obtained for PulD 28 -42/259 -660 and PulD 28 -42/259 -660 T470I (0.14 Ϯ 0.04 and 0.07 Ϯ 0.05 min Ϫ1 , respectively), the rate constant is not easily determined for PulD 28 -42/259 -660 I322S, which remains largely monomeric upon ZW 3-14 treatment in kinetic experiments (Fig. 7, inset, and supplemental Fig. S5, D-F). However, PulD 28 -42/259 -660 I322S monomers decayed with a rate constant of 0.34 Ϯ 0.14 min Ϫ1 (Fig. 7, inset), providing an estimate for the multimerization rate constant. This places the rate constants for all PulD 28 -42/259 -660 variants within 2-fold of each other, indicating that E a would change by less than 2 kJ/mol/multimer. The effect of the I322S substitution on SDS-resistant prepore stability was also estimated via the ⌬⌬G 0 ϭ 8.33 kJ/mol between N 3 and N 3 I322S. The sharp reduction in the amount of mixed multimers formed with PulD fl WT when the number of PulD 28 -42/259 -660 I322S or PulD 28 -42/259 -660 I322S/T470I protomers exceeded six (Fig. 6B), suggested that the maximal destabilization tolerated in the N 3 -ring to produce a large amount of native multimers is ϳ50 kJ/mol (Ϸ6 ϫ 8.33 kJ/mol). If this would be attributed to an increase in E a , the mixed multimers would take years to fold (because k unaltered /k mixed ϭ exp(Ϫ⌬E a /RT)ork T470I /k mixed ϭexp(Ϫ⌬E a /RT)).Thisisinconsistent with the abundant SDS and urea-resistant PulD-mixed multimers observed within 6 h. Thus, kinetic and thermodynamic measurements suggested that the I322S substitution must primarily destabilize the SDS-resistant prepore.

Discussion
Although each PulD protomer only shares parts of the C domain to form the dodecameric transmembrane pore and the gate that closes it (6), PulD will not assemble into such a struc- ture in the absence of N 3 (15). The results presented here reveal why: N 3 provides thermodynamic stability to the obligate prepore that is formed prior to PulD membrane insertion. Substitution of Ile 322 into Ser in N 3 increases the conformational dynamics of structural elements in N 3 , in particular in the ␣-helix that is next to the C domain and in the ␤-sheet opposite of this helix. Essential interactions between neighboring N 3 domains and/or between N 3 and the C domain might be disrupted in the context of the PulD dodecamer, but mixed multimers suggest that the global stability in the N 3 -ring predominates. The ⌬⌬G 0 ϭ 4.69 kJ/mol between N 3 I322S and N 3 ⌬L 2 I322S indicates that decreasing the ⌬G 0 of the multimer with ϳ56 kJ/mol (ϭ12 ϫ 4.69 kJ/mol) prevents the formation of the native PulD structure. The same idea emerges from the ⌬⌬G 0 ϭ 8.33 kJ/mol between N 3 WT and N 3 I322S and the observation that six PulD fl WT protomers are required to form a mixed multimer with PulD 28 -42/259 -660 I322S (6 ϫ 8.33 kJ/mol ϭ 50 kJ/mol). This is the equivalent of few van der Waals contacts or a H-bond per monomer. We propose that the N 3 -ring forms a seal onto the assembled C domains and must attain a critical stability to facilitate the conformational rearrangements in the C domain that drive pore formation (13), while preventing multimer disruption.
Insights into the folding of OMPs that multimerize into a shared transmembrane pore mainly come from the isolation of intermediates, but the kinetic and thermodynamic relationship with the native structure is mostly unknown (14,(23)(24)(25). Previous reports (13,14,26) and data obtained here help to define these relationships for PulD by characterizing intermediates that form in the presence of a lipid membrane in detail after detergent solubilization (Fig. 7). Combined, these data indicate that four structures reside in local energy minima along the reaction coordinate: the monomer (Mo), an SDS-sensitive early prepore (EP), the sealed SDS-resistant late prepore (LP), and the pore or native multimer (NMu) (Fig. 7). The largest driving force for PulD pore formation likely comes from membrane insertion of the transmembrane domain, consistent with the remarkable stability of NMu and the ease by which LP is dissociated by moderate urea concentrations (14). CD spectra and EM reconstructions support that PulD traverses the membrane by a ␤-barrel that is 9 nm in diameter (6,13,15). To create such a ␤-barrel, each PulD protomer should contribute at least 4 ␤-strands (27). With each ␤-strand contributing 10 kJ/mol to OMP stability (28), this amounts to an overall folding free energy of at least 480 kJ/mol (Fig. 7). Yet, the balance between EP and LP is attenuated by a modest ⌬⌬G 0 of 50 kJ/mol/dodecamer. Furthermore, the low multimerization efficiency of PulD 28 -42/259 -660 I322S, as determined by SDS-PAGE, suggests that the ⌬⌬G 0 associated with the Mo to EP transition is of the same order of magnitude (ϭ12 ϫ 8.33 kJ/mol; Fig. 7).
Can this reflect the in vivo pathway? PulD belongs to an expanding group of BAM-independent OMPs that rely on lipid-assisted self-assembly once monomers are delivered to the OM via the Lol system, which in the case of PulD is mediated by the lipoprotein PulS (2,3,12,26). This provides a rationale to study the membrane-associated steps of PulD assembly in a controlled in vitro environment in the presence of liposomes and to compare the results from in vitro studies to the in vivo pathway. Whereas PulS targets PulD to the OM in vivo, PulS accelerates PulD multimerization in vitro (26), providing a first driving force for PulD folding. The binding energy between PulD S domains and PulS was determined to be 40 -50 kJ/mol (10). With the Mo to EP and EP to LP transitions each stabilizing the PulD multimer 50 kJ/mol (Fig. 7), the formation of these structures does not provide the energy to dissociate PulD/PulS heterodimers. Interestingly, dissociation is not required to form NMu in vivo (26) and PulS co-purifies with PulD from in vivo sources (29). PulS prevents proteolysis of PulD protomers in vivo (30), which could rationalize the necessity for a prolonged PulD/PulS association as PulD protomers join and break free from growing complexes until the correct conformation is found. Alternatively, the PulD/PulS heterodimer is required to retain PulD folding competence during multimerization. Hence, whereas the rough free energy landscape from Mo to EP only provides modest stability, it effectively lowers the entropy associated with PulD multimerization, providing a second driving force for PulD assembly. Lowering entropy ultimately facilitates rapid lipid-assisted membrane insertion, a third driving force in PulD assembly. In this context it is worth noting that a loss of 100 kJ/mol in a PulD 28 -42/259 -660 I322S dodecamer relative to PulD 28 -42/259 -660 is equivalent to the folding free energy of a small to medium-sized BAM-dependent OMP.
Multimeric OMPs (3,14,(23)(24)(25)31) and bacterial poreforming toxins and eukaryotic-pore forming complexes (32) that form prepores typically contain at least one domain in the membrane periphery. Folding pathways are not easily obtained, because of the fast assembly rates and the complex environment that these proteins often require. However, multiple sequential pre-integration structures have been described for the bacterial toxins aerolysin (33) and perfringolysin O (34). A salt bridge between neighboring domains in the membrane periphery critically stabilizes an EP into an LP before NMu formation occurs in perfringolysin O, but removing it increases E a only by 2 kJ/mol/monomer (34). Because a salt bridge is typically stronger, it is possible that it also acts to seal EP into LP. In conclusion, we propose that the energies associated with EP-to-LP transitions driven by domains in the membrane periphery will emerge as key driving forces for membrane insertion of all constitutive and non-constitutive multimeric complexes that assemble independently of a membrane-embedded insertase.

Experimental Procedures
Strains and Growth Conditions-Cloning and stress response experiments were performed in the Escherichia coli strain K-12 PAP105 (⌬(lac-pro) FЈ (lacI q1 ⌬lacZM15 proAB ϩ Tn10)). E. coli strain MC4100 PAP7447 (FЈ lacI q pro ϩ Tn10)], with pulS, pulA, and pulC-O integrated into malPp and with a large deletion in pulD, was used for in vivo secretion (11). Strains were grown at 30°C in LB medium with 100 g/ml of ampicillin and/or 25 g/ml of chloramphenicol (as appropriate). Expression of the pul genes was induced with 0.4% maltose.
Plasmid Construction and Site-directed Mutagenesis-Plasmids and primers used are given in supplemental Table S2 Step 2 involved mixing the purified PCR products at equimolar ratios to generate a DNA fragment that encoded for the entire PulD 28 -42/124 -190/342-660 amino acid sequence between the SphI and EcoRI restriction sites using primers ING318 and ING323. In step 3 this fragment and pIVEX2.3 MCS were digested with the restriction enzymes SphI and EcoRI and ligated to yield pCHAP3359. The same procedure was used to generate pCHAP3357 that encoded for PulD 28 -42/188 -260/342-660 , in which N 3 was replaced by N 2 . The  primer pairs ING318/ING324, ING325/ING326, ING327/  ING323, and ING318/ING323 were used in PCR in step 1a, step 1b, step 1c, and step 2, respectively. pCHAP3358 that encodes PulD fl I322S was created by digesting A6 with AgeI and EcoRI and the short fragment carrying the mutation encoding for I322S was ligated into the long fragment of pCHAP3731 digested with the same enzymes. Plasmids encoding for PulD fl L168S (pCHAP3355) and PulD fl L241S (pCHAP3356) were obtained by site-directed mutagenesis of pCHAP3731 using the primer pairs L168SF/R and L241SF/R, respectively. The plasmid encoding PulD 28 -42/188 -260/342-660 L241S (pCHAP3357) was obtained by site-directed mutagenesis of pCHAP3375 using the primers L241SF/R. To delete the sequence encoding for L 2 (amino acids 205 to 308) in N 3 , the purified PCR product of pCHAP3716 or A6 and the phosphorylated primers ING349/ING350 were ligated to yield pCHAP3369 (encoding for PulD 28 -42/259 -660 ⌬L 2 ) or pCHAP3374 (encoding for PulD 28 -42/259 -660 ⌬L 2 I322S), respectively. pCHAP3395 and pCHAP3396, encoding PulD fl ⌬L 2 and PulD fl ⌬L 2 I322S, respectively, were obtained by ligating the short fragments from pCHAP3369 and pCHAP3374 digested with AgeI and EcoRI (encoding for the L 2 -deletion and the I322S substitution, respectively) into pCHAP3731 digested with the same enzymes. The plasmid encoding PulD 28 -42/259 -660 I322S/T470I (pCHAP3328) was obtained by exchanging the short fragment generated by AgeI and AflII digestion of pCHAP3727 (which contains the sequence for the T470I substitution) with that generated from A6 after digestion with the same enzymes (in which the large fragment contains the sequence with the I322S substitution). The sequences encoding for N 3 WT, N 3 I322S, N 3 ⌬L 2 , and N 3 ⌬L 2 I322S were obtained by PCR from the pCHAP3716, A6, pCHAP3369, and pCHAP3374 using primers ING347 and ING348. Digestion of the amplicons with NdeI and XmaI enabled ligation in-frame with the His 6 tag encoded on pIVEX2.3 MCS to give pCHAP3362, pCHAP3368, pCHAP3392, and pCHAP3393, respectively. To generate the pCHAP3378 that carries the pulD fl I322S gene with the signal sequence and the His 6 tag, the pulD fl I322S was excised from A6 using AgeI and BlpI and inserted into pCHAP3671 digested with the same enzymes. To insert pulD fl I322S in a moderate copy number vector, pCHAP3378 was digested with EcoRI and HindIII and the resulting fragment carrying pulD fl I322S was inserted into pSU18 digested with the same enzymes to yield pCHAP3389. Finally, as pCHAP3389 encodes for a PulD fl I322S variant with a His 6 tag, pCHAP3389 and pCHAP3635 were digested with EcoRI and HpaI and the fragment carrying pulD fl I322S was ligated into pCHAP3635 to yield pCHAP3394. The same procedure was followed to generate plasmids encoding for PulD fl ⌬L 2 and PulD fl ⌬L 2 I322S, using pCHAP3369 and pCHAP3374 as parent plasmids to give pCHAP3390 and pCHAP3391, respectively.
Gel Electrophoresis and Immunoblotting-Proteins were separated by electrophoresis on a 4 -10% SDS-polyacrylamide gel. Samples were mixed with SDS loading buffer (final concentra-tion of 62.5 mM Tris-HCl, pH 6.8, 12.5% glycerol, 2% SDS). Following SDS-PAGE, the proteins were transferred onto a nitrocellulose sheet by semi-dry blotting. PulD variants, PspA and OmpF were detected using anti-PulD, anti-PspA, and anti-OmpF antibodies, respectively. Primary antibodies were detected by a secondary antibody coupled to horseradish peroxidase (GE Healthcare). Peroxidase activity was measured by the chemiluminescence produced by ECL2 (Pierce) and integrated on a Typhoon imager (GE Healthcare).
PspA Induction and Pullulanase Secretion-PspA induction was performed in the absence or presence of PulS (pCHAP585). Cultures were inoculated from overnight growths at D 600 ϭ 0.15 and grown for an additional 6 h at 30°C before cell harvesting. PspA production was analyzed by SDS-PAGE and immunoblotting of cells in SDS-loading buffer to a final concentration of 10 D 600 /ml.
The same growth conditions were used to perform the secretion assay. Cells were grown to a D 600 ϭ 1.0 -1.5 and chilled on ice. The pullulanase assay was performed as described previously (35). The level of secretion is represented as the percentage of the total amount of pullulanase measured in unlysed cells.
In Vitro PulD Synthesis, Zwittergent 3-14 Solubilization, and Mixed Multimer Formation-In vitro PulD synthesis was done with the RTS100 E. coli HY kit (18). The reaction mixture was supplemented with lecithin liposomes at a final concentration of 2 mg/ml and incubated for 6 h at 30°C. For ZW 3-14 treatment of PulD variants, part of the reaction mixture was mixed with a final concentration of 1.5% ZW 3-14 in 25 mM Tris, pH 7.2, and incubated for 1 h at room temperature. Controls were diluted with detergent-free buffer. To assess the effect of urea or DDM on the multimeric state of PulD 28 -42/259 -660 I322S, liposomes, isolated from the in vitro synthesis reaction (16,100 ϫ g for 15 min), were resuspended into 100 mM Tris, pH 7. 5, 500 mM NaCl and diluted 2-fold with 1, 2, 4, 6, or 8 M urea or 2% DDM. After incubation for 1 h at room temperature, a final concentration of 1.5% ZW 3-14 (and 25 mM Tris, pH 7.5, 125 mM NaCl) was added to all samples, except the controls, and incubated overnight. PulD multimers were dissociated by phenol where indicated (11). Formation of mixed multimers was achieved by supplementing the reaction mixture with 10 mg/ml of DNA in a 1:3 ratio of the plasmids carrying the pulD fl and pulD 28 -42/259 -660 genes, respectively, in the presence of lecithin liposomes. Liposomes were sedimented, resuspended, and divided into two parts. Liposomes were diluted in 25 mM Tris, pH 7.2, with or without 4 M urea (final) and incubated 1 h at room temperature. All samples were analyzed by SDS-PAGE and immunoblotting.
Folding Kinetics Followed by SDS Treatment-Folding kinetics of PulD 28 -42/259 -660 variants were obtained by mixing aliquots of the synthesis reaction at the time points indicated with SDS-loading buffer to stop folding and incubated on ice before analysis by SDS-PAGE (13). Where kinetics relied solely on integration of the monomer, curves were corrected for linear degradation of the monomer. Curves were fitted to a single exponential equation in Origin 8.0.
Equilibrium Unfolding-N 3 variants were purified as described before (10), diluted to 0.2 mg/ml into 50 mM sodium phosphate, pH 8.0, and the appropriate amount of urea and incubated overnight to reach equilibrium. CD spectra were taken at 25°C on an AVIV spectrometer between 190 and 260 nm (where possible) at a rate of 20 nm/min and a bandwidth of 1 nm in a 0.1-cm cuvette. Spectra were averaged over three measurements. For equilibrium unfolding experiments, 100 data points taken at 220 nm were averaged. The data were fitted to y ϭ ((y N ϩ m Transmission Electron Microscopy (TEM) and Image Processing-His-tagged PulD 28 -42/259 -660 variants were extracted from liposomes by 3% ZW 3-14 in 50 mM Tris, pH 7.5, and 250 mM NaCl and purified using a cobalt affinity chromatography (Talon, Clontech). Solubilized PulD 28 -42/259 -660 variants were bound to the resin for 1 h and washed with 5 column volumes of 50 mM Tris, pH 7.5, 250 mM NaCl, and 0.6% ZW 3-14 before elution in the same buffer supplemented with 5 mM EDTA. Eluted PulD 28 -42/259 -660 variants were concentrated and frozen for EM analysis. His-tagged proteins were used as they increased the amount of particles that adsorbed perpendicular on the EM grid.
A 4-l aliquot was adsorbed onto a glow discharged carbon film-coated copper EM-grid, washed with three droplets of pure water, and subsequently negatively stained with 2% (w/v) uranyl-acetate. The grids were imaged using a Philips CM10 TEM (FEI Company, Eindhoven, the Netherlands) operating at 80 kV. The images were recorded by the 2k ϫ 2k side-mounted Veleta CCD camera (Olympus, Germany) at a magnification of 130,000. The pixel size at the sample level was 3.7 Å.
Image processing was done in the EMAN2 software package (36). The images were contrast transfer function corrected and particle projections were semi-automatically selected. e2refine2d was used to classify the particle projections and yielded reference-free class averages from a population of mixed, unaligned particle projections. The representative class averages with the best signal-to-noise ratio were selected and gathered in a gallery.
Sample Preparation for H/DX-MS-All samples were prepared in triplicate using a fully automated LEAP H/DX Pal robot (LEAP Technologies, Carrboro, NC). Prior to deuterium labeling, N 3 variants (15 M in 50 mM HEPES buffer, pH 7.0) were equilibrated for 60 min at room temperature and placed at 4°C in the protein compartment of the robot. H/DX was initiated by diluting 2 l of each protein (30 pmol) with 48 l of HEPES buffer made with 99.9% D 2 O, pD 7.0, and at 20°C. At time points between 10 s and 4 h, 30 l of the labeling reaction was removed, quenched with 60 l of ice-cold 1.5% formic acid to lower the pH to 2.5, and immediately analyzed by MS. Fully deuterated samples were prepared manually by 24 h incubation at 60°C (final D 2 O content of 96%), quenched as described above and frozen until LC-MS analysis.
H/DX-MS Data Acquisition-Quenched samples (10 pmol) were injected into a refrigerated Waters nanoACQUITY UPLC HDX system maintained at 0°C (37). For global H/DX-analyses, samples were desalted for 2 min on a C4 trap column (Van-