Type III secretion system effector proteins are mechanically labile

Multiple Gram-negative bacteria encode Type III secretion systems (T3SS) that allow them to inject effector proteins directly into host cells to facilitate colonization. To be secreted, effector proteins must be at least partially unfolded to pass through the narrow needle-like channel (diameter < 2 nm) of the T3SS. Fusion of effector proteins to tightly packed proteins—such as GFP, ubiquitin, or dihydrofolate reductase (DHFR)—impairs secretion and results in obstruction of the T3SS. Prior observation that unfolding can become rate limiting for secretion has led to the model that T3SS effector proteins have low thermodynamic stability, facilitating their secretion. Here, we first show that the unfolding free energy (ΔG0unfold) of two Salmonella effector proteins, SptP and SopE2, are 6.9 and 6.0 kcal/mol respectively, typical for globular proteins and similar to published ΔG0unfold for GFP, ubiquitin, and DHFR. Next, we mechanically unfolded individual SptP and SopE2 molecules by AFM-based force spectroscopy. SptP and SopE2 unfolded at low force (Funfold ≤ 17 pN @ 100 nm/s), making them among the most mechanically labile proteins studied to date by AFM. Moreover, their mechanical compliance is large, as measured by the distance to the transition state (Δx‡ = 1.6 and 1.5 nm for SptP and SopE2, respectively). In contrast, prior measurements of GFP, ubiquitin, and DHFR show them to be mechanically robust (Funfold > 80 pN) and brittle (Δx‡ < 0.4 nm). These results suggest that effector protein unfolding by T3SS is a mechanical process and that mechanical lability facilitates efficient effector protein secretion.

Methods
Circular dichroism data was collected and analyzed for Fig. 1 as follows:

Protein was removed from the -80 ºC freezer, thawed, and then centrifuged at 21,000 rcf for 5 min. A 10 M urea solution was deionized using BioRad AG 501-X8 resin (50 g beads/L urea) for 1 h and vacuum filtered through a 0.22-μm membrane to remove the resin. Urea concentration was measured using an Abbe refractometer. For equilibrium unfolding measurements, serial dilutions of urea with a fixed protein concentration of 0.05 mg/ml  as well as corresponding no-protein blanks were prepared  by mixing 10X SptP buffer (100 mM Tris base, pH 8.0, 1.5 M sodium sulfate, and 5 mM TCEP) or 10X SopE2 buffer (250 mM HEPES, pH 7.2, 1.5 M NaCl, and 5 mM TCEP) with ultrapure water, 10 M urea stock, and protein to 1X buffer concentration. Samples were incubated in a 25 ºC water bath for 1–3 days to reach equilibrium before CD spectra were collected.

Measurements were performed using a quartz cuvette (Hellma) with a 1-mm path length on an Applied PhotoPhysics ChiraScan Plus spectrophotometer. Measurement parameters: λ = 212.5–260 nm; step size = 0.5 nm; bandwidth = 1.0 nm; time per point = 0.5 s; and 3 repeats. The instrument was thoroughly purged with nitrogen to prevent ozone formation. Temperature was held at 25 °C with a Peltier sample holder and the temperature recorded using the temperature probe. Prior to loading, samples were spun at 18,000 rcf for 5 min. We measured a control sample as “blank” before every protein sample. Following this pair of measurements, the cuvette was serially rinsed with several mL each of 10 M urea, urea free buffer, 1% cleaning solution (Hellmanex), and ultrapure water. The cuvette was then filled with ultrapure water and a CD spectra taken to ensure no protein adhered to the cuvette. The cuvette was then rinsed with absolute ethanol and dried using filtered house air. This was repeated for every concentration of urea.

We analyzed the CD data using Applied Photophysics software. First, the three independent measurements were averaged. The subsequent spectrum was smoothed using the Savitzky–Golay algorithm with a window size of 12 points. This smoothing was done on both the protein-containing sample and the blank. We then subtracted the smoothed blank spectrum from the smoothed protein-containing spectrum to give the final, baseline corrected spectrum. After this analysis was done for all urea concentrations, the ellipicity at λ = 222 nm was plotted as a function of urea concentration. This plot was fit with an equation to determine the free-energy of unfolding assuming a two state system which accounts for a sloping baseline.

AFM data was collected and analyzed as follows for Fig. 2:

AFM experiments were performed on a Cypher ES (Asylum Research) in a temperature-controlled closed fluidic cell (T = 25 ºC). The stiffness (k) of the FIB-modified cantilevers was calibrated using the thermal method (81) far from the surface while sensitivity was measured by pressing the cantilever into hard contact with the surface. The cantilevers had an average k ≈ 6.5 pN/nm. Force-extension curve acquisition was initiated by pressing the cantilever into the surface at 100 pN for 0–200 ms depending on the surface polyprotein concentration. This comparatively low indentation force was enabled by our site-specific, cohesin-dockerin-based coupling between the tip and the polyprotein. To minimize the compliance of the polyprotein construct, we used only a single marker domain and short PEG linkers (MW = 600 D), which facilitated detecting proteins that unfold at low force and low extension [Fig. 2B,C (inset)]. We retracted the cantilever at 100–3,200 nm/s while digitizing at 50 kHz. We acquired multiple traces per sample by probing the surface in a raster scan, moving the AFM tip in a grid pattern with each location separated by 150 nm. Each spot was probed 10 times unless a molecule was detected, in which case the spot was continually sampled until ~20 consecutive attempts failed to yield a connection. This meant that an individual protein could be repeatedly probed. We found that both SopE2CD and SptPCD refolded well, and repeated cycles of unfolding and refolding did not affect the observed unfolding forces (SI Appendix, Fig. S3). The high-bandwidth records were boxcar averaged to the indicated bandwidths for analysis and presentation (1–5 kHz). Force was determined by cantilever deflection accounting for the sensitivity and stiffness of each cantilever. Extension was calculated from the movement of the sample stage minus the deflection of the cantilever. The loading rate (pN/s) for each unfolding event in a force−extension curve was calculated by fitting a line to the force-versus-time curve immediately preceding effector protein unfolding. For the effector protein unfolding-force analysis, only the first unfolding event was used when an unfolding intermediate was observed. A small percentage for the force-extension curves showed atypically high unfolding forces for the initial unfolding of SptPCD SopE2CD (8 and 2% respectively). These records were excluded from analysis as they most likely represented rare tip-sample surface adhesion and/or unfolding of a misfolded protein.

The data for Fig. S1 was collected as follows:

To demonstrate reversibility of urea denaturation, we refolded urea denatured SopE2CD and SptPCD by dilution. SopE2CD was buffered with 25 mM HEPES, pH 7.2, 150 mM NaCl, and 0.5 mM TCEP while SptPCD was buffered with 10 mM Tris, pH 8.0, 150 mM sodium sulfate, and 0.5 mM TCEP. Three sample conditions were used, a high urea control sample [4.7 M (SopE2CD); 5.6 M (SptPCD)], low urea control sample [0.38 M (SopE2CD); 0.94 M (SptPCD)], and refolded sample (diluted from 4.7 M to 0.38 M for SopE2CD and from 5.6 M to 0.94 M for SptPCD). Urea buffer without protein was also prepared as a blank. The purpose of the high and low urea controls was to show that the protein denatured under the experimental conditions and to give a signal for native protein for comparison. Samples were prepared and allowed to equilibrate for one day in a 25 ºC water bath. For refolding, the equilibrated samples were added dropwise to a glass vial with a stir bar containing either buffer containing the same amount of urea as the sample (for controls) or buffer with no urea (for the refolding sample). CD spectra were obtained as described in Circular dichroism measurement and analysis in the main text. Two independent high and low urea concentration samples were measured and three independent refolded samples were measured (Fig. S1).

The data for Fig. S3 were collected as follows:

Comparisons of the mean Funfold of the first unfolding event of SptP or SopE2 to the mean Funfold of the refolded protein. Measurements were collected with a pulling velocity of 1600 nm/s [Nfirst = 41; Nrefold = 12 (SptP); Nfirst = 18; Nrefold = 27 (SopE2)]. Error bars represent the SEM. In the case of an unfolding intermediate, only the first unfolding event of the effector protein was analyzed.

多种革兰氏阴性菌（Gram-negative bacteria）编码Ⅲ型分泌系统（Type III secretion systems, T3SS），该系统可使其将效应蛋白直接注射至宿主细胞内以促进定殖。效应蛋白要被分泌，必须至少部分解折叠，才能通过T3SS直径小于2 nm的针状狭窄通道。将效应蛋白与紧密折叠的蛋白——如绿色荧光蛋白（GFP）、泛素（ubiquitin）或二氢叶酸还原酶（dihydrofolate reductase, DHFR）——融合，会损害分泌过程并造成T3SS阻塞。此前有研究观察到，解折叠过程可能成为分泌的速率限制步骤，由此提出假说：T3SS效应蛋白具有较低的热力学稳定性，以促进其分泌。

本研究首先证实，两种沙门氏菌效应蛋白SptP和SopE2的解折叠自由能（ΔG⁰_unfold）分别为6.9 kcal/mol和6.0 kcal/mol，这一数值符合球状蛋白的典型特征，且与已发表的GFP、泛素及DHFR的ΔG⁰_unfold相近。随后，我们通过基于原子力显微镜的力谱（AFM-based force spectroscopy）技术对单个SptP和SopE2分子进行机械解折叠实验。结果显示，SptP和SopE2在低力下即可解折叠（100 nm/s条件下，解折叠力Funfold ≤17 pN），是目前通过原子力显微镜（AFM）研究的力学稳定性最差的蛋白之一。此外，它们的力学柔顺性较强，通过到达过渡态的距离（Δx‡：SptP为1.6 nm，SopE2为1.5 nm）可测得这一特性。与之形成对比的是，此前针对GFP、泛素及DHFR的测量结果显示，这些蛋白力学稳定性较强（Funfold >80 pN）且脆性较高（Δx‡ <0.4 nm）。上述结果表明，T3SS介导的效应蛋白解折叠属于力学过程，而蛋白的力学不稳定性可促进效应蛋白的高效分泌。

## 材料与方法
### 图1的圆二色性（Circular dichroism, CD）数据收集与分析
如下所述：
1. 样品前处理：将保存在-80 ℃冰箱中的蛋白取出解冻，随后以21000 rcf离心5 min。使用BioRad AG 501-X8树脂（50 g树脂/L尿素）对10 M尿素溶液进行1 h脱离子处理，随后通过0.22 μm滤膜真空过滤以去除树脂。使用阿贝折射仪测定尿素浓度。
2. 平衡解折叠样品制备：将10× SptP缓冲液（100 mM Tris碱，pH 8.0，1.5 M硫酸钠，5 mM TCEP）或10× SopE2缓冲液（250 mM HEPES，pH 7.2，1.5 M氯化钠，5 mM TCEP）与超纯水、10 M尿素原液及蛋白混合，配置得到固定蛋白浓度为0.05 mg/ml的系列尿素稀释样品，以及对应的无蛋白空白对照。将样品置于25 ℃水浴中孵育1~3天以达到平衡，随后收集CD光谱。

使用光程为1 mm的Hellma石英比色皿，在Applied PhotoPhysics ChiraScan Plus分光光度计上进行测量。测量参数如下：波长范围212.5~260 nm，步长0.5 nm，带宽1.0 nm，每个数据点采集时间0.5 s，重复测量3次。仪器全程用氮气吹扫以防止臭氧生成。使用珀尔帖样品池将温度维持在25 ℃，并通过温度探头记录实际温度。上样前，样品以18000 rcf离心5 min。每测量一个蛋白样品前，均先测量空白对照样品。完成一组样品测量后，依次用数毫升10 M尿素、无尿素缓冲液、1% Hellmanex清洁液及超纯水对比色皿进行连续冲洗，随后向比色皿中加入超纯水并收集CD光谱，以确认无蛋白附着在比色皿内壁。之后用无水乙醇冲洗比色皿，并用过滤后的室内空气吹干。该冲洗流程需针对每个尿素浓度的样品重复执行。

使用Applied Photophysics配套软件对CD数据进行分析。首先对3次独立测量结果取平均值，随后使用Savitzky-Golay算法（窗口大小为12个数据点）对光谱进行平滑处理，含蛋白样品与空白对照均需执行该平滑步骤。将平滑后的空白光谱从平滑后的含蛋白光谱中减去，得到最终的基线校正光谱。对所有尿素浓度的样品完成该分析后，以波长222 nm处的椭圆率为纵坐标，尿素浓度为横坐标绘图，并采用考虑倾斜基线的两态模型拟合曲线，以计算解折叠自由能。

### 图2的原子力显微镜（AFM）数据收集与分析
如下所述：
AFM实验在Cypher ES（Asylum Research）上完成，实验环境为温度可控的封闭式流体池（温度维持25 ℃）。使用热校准法对聚焦离子束（FIB）修饰的悬臂梁刚度（k）进行校准，校准过程远离样品表面；悬臂梁灵敏度则通过将其压至样品表面硬接触进行测定。实验所用悬臂梁的平均刚度约为6.5 pN/nm。
力-拉伸曲线采集流程：将悬臂梁以100 pN的力压入样品表面，压入时间0~200 ms，具体时长取决于表面多聚蛋白的浓度。该低压入力得益于我们采用的基于黏连蛋白-dockerin的位点特异性偶联方式，实现了针尖与多聚蛋白的连接。为降低多聚蛋白构建体的柔顺性，我们仅使用单个标记结构域与短PEG连接子（分子量600 D），这一设计便于检测在低力与低伸长量下解折叠的蛋白（参见图2B、C插图）。以100~3200 nm/s的速度收回悬臂梁，同时以50 kHz的采样率进行数字化采集。通过光栅扫描方式对样品表面进行多点探测，每个探测位置间距为150 nm，以此获取多个样品的测量轨迹。每个位置默认探测10次，若检测到蛋白分子，则持续对该位置进行采样，直至连续约20次尝试均未获取到蛋白连接信号为止，这使得单个蛋白分子可被多次探测。
我们发现SptP_CD与SopE2_CD均具有良好的复性能力，且重复的解折叠-复性循环不会影响观测到的解折叠力（参见补充材料图S3）。将高带宽的记录数据进行箱式平均，以得到用于分析与展示的带宽范围（1~5 kHz）。力的计算基于悬臂梁挠度，并结合每个悬臂梁的灵敏度与刚度参数；伸长量则通过样品台移动距离减去悬臂梁挠度得到。针对力-拉伸曲线中的每个解折叠事件，通过对解折叠前即刻的力-时间曲线进行线性拟合，计算其加载速率（pN/s）。在进行效应蛋白解折叠力分析时，若观测到解折叠中间体，则仅选取第一个解折叠事件进行分析。少量力-拉伸曲线显示，SptP_CD与SopE2_CD的初始解折叠过程出现了异常高的解折叠力（分别占比8%和2%），此类记录被排除在分析之外，因为它们大概率代表了罕见的针尖-样品表面粘附，或是错误折叠蛋白的解折叠事件。

### 图S1的数据收集
为验证尿素变性的可逆性，我们通过稀释法对尿素变性的SopE2_CD与SptP_CD进行复性。SopE2_CD所用缓冲液为25 mM HEPES，pH 7.2，150 mM氯化钠，0.5 mM TCEP；SptP_CD所用缓冲液为10 mM Tris，pH 8.0，150 mM硫酸钠，0.5 mM TCEP。设置三组样品条件：高尿素对照组[4.7 M（SopE2_CD）；5.6 M（SptP_CD）]、低尿素对照组[0.38 M（SopE2_CD）；0.94 M（SptP_CD）]以及复性样品组（SopE2_CD从4.7 M稀释至0.38 M，SptP_CD从5.6 M稀释至0.94 M）。同时制备无蛋白的尿素缓冲液作为空白对照。高、低尿素对照组的设置用于验证蛋白在实验条件下发生了变性，并为天然蛋白提供对照信号。样品制备完成后置于25 ℃水浴中平衡1天。复性操作时，将平衡后的样品逐滴加入带有磁力搅拌子的玻璃小瓶中，小瓶内分别装有与样品尿素浓度相同的缓冲液（对照组）或无尿素缓冲液（复性样品组）。CD光谱的获取方法与正文中圆二色性测量及分析流程一致。本实验共测量2组独立的高、低尿素浓度样品，以及3组独立的复性样品（参见补充材料图S1）。

### 图S3的数据收集
对比SptP或SopE2首次解折叠事件的平均解折叠力，与复性蛋白的平均解折叠力。测量时拉伸速度为1600 nm/s[SptP：首次解折叠样本量N_first=41，复性样本量N_refold=12；SopE2：首次解折叠样本量N_first=18，复性样本量N_refold=27]。误差棒代表标准误（standard error of the mean, SEM）。若观测到解折叠中间体，则仅分析效应蛋白的首次解折叠事件。