Efficient cellular solid-state NMR of membrane proteins by targeted protein labeling
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Solid-state NMR spectroscopy (ssNMR) has made significant progress towards the study of membrane proteins in their native cellular membranes. However, reduced spectroscopic sensitivity and high background signal levels can complicate these experiments. Here, we describe a method for ssNMR to specifically label a single protein by repressing endogenous protein expression with rifampicin. Our results demonstrate that treatment of E. coli with rifampicin during induction of recombinant membrane protein expression reduces background signals for different expression levels and improves sensitivity in cellular membrane samples. Further, the method reduces the amount of time and resources needed to produce membrane protein samples, enabling new strategies for studying challenging membrane proteins by ssNMR.
KeywordsSolid-state NMR Membrane protein Cellular membranes Protein expression
Membrane proteins (MPs) are often challenging to study by conventional structural biology methods and their native environment, a cellular membrane, is very heterogeneous and difficult to replicate in vitro. Although many membrane mimetics are available, the choice of mimetic can alter the biophysical properties of MPs (see, for example, Zhou and Cross 2013; Etzkorn et al. 2013). This phenomenon is one reason for the recent push towards crystallizing MPs in lipidic mesophases (Caffrey et al. 2012, for a review).
Solid-state nuclear magnetic resonance spectroscopy (ssNMR) is one method available to study the structure and dynamics of MPs in a lipid bilayer. ssNMR has been used successfully to determine the structure of membrane proteins (see, for example, Andronesi et al. 2005; Das et al. 2012; Wang et al. 2013a; van der Cruijsen et al. 2013), as well as investigating molecular interactions (Miao and Cross 2013, for a review) and function (Baker and Baldus 2014, for a review). However, most of these studies have used synthetic lipid bilayers, which can mimic the physical environment of cellular membranes but often lack their chemical heterogeneity. In addition to more closely resembling the cellular environment, the use of native membranes can remove the need to purify the MP of interest, eliminating any potential for structural disruptions during the solubilization process and speeding up sample preparation.
Several groups have demonstrated that ssNMR can be used to study structure and dynamics of proteins embedded in native lipid membranes (Etzkorn et al. 2007; Fu et al. 2011; Jacso et al. 2012; Miao et al. 2012; Kulminskaya et al. 2012; Ward et al. 2015). Previous work has demonstrated that it is possible to study MPs in cellular envelopes and whole cells of Escherichia coli by conventional (Renault et al. 2012b; Kaplan et al. 2015) and dynamic nuclear polarization (DNP)-enhanced ssNMR (Renault et al. 2012a; Yamamoto et al. 2014; Kaplan et al. 2015). In particular, we could show that studies using cellular envelopes or whole cells of Gram-negative bacteria are facilitated by use of a special E. coli deletion strain, to remove signals from naturally highly abundant outer MPs, i.e., OmpA and OmpF. In these preparations, signals from other cellular components, such as lipids, nucleotides, peptidoglycan and lipopolysaccharides, remain visible at intensities similar to that of the (overexpressed) protein of interest (Baldus 2015). Such non-proteinaceous correlations can help to refine the supramolecular structure of a protein in a membrane setting (see, e.g., Weingarth and Baldus 2013). Further, these signals can be used to study the structure of other molecular components such as RNA (Renault et al. 2012a) or material in the cell walls of bacteria and plants (Dick-Pérez et al. 2011, 2012; Wang et al. 2012, 2014), as well as helping to clarify the cellular distribution of added reagents, such as radicals for DNP experiments (Takahashi et al. 2013).
On the other hand, these cellular signals can complicate the analysis of spectra of MPs that have not previously been characterized in synthetic bilayers. To reduce the challenges posed by the high background levels in cellular membrane samples, it would be beneficial to have a system by which isotope labels could be selectively incorporated into a MP of interest. Here, we demonstrate that treatment of cells with the antibiotic rifampicin, to inhibit the native RNA polymerase, produces native MP samples with significantly less background in ssNMR spectra. Previously, this concept was used for preparation of labeled proteins for solution-state NMR without purification (Almeida et al. 2001) with potential benefits to prevent cell lysis in the context of in-cell solution-state NMR experiments (Cruzeiro-Silva et al. 2006). During rifampicin treatment, the E. coli RNA polymerases are inhibited, preventing endogenous gene expression. However, systems for over-expression of proteins in E. coli, such as the T7 system, often use non-E. coli RNA polymerases that are not inhibited by rifampicin. If one induces expression of the heterologous protein after rifampicin treatment, the heterologous protein should be produced while endogenous protein production should decline. The suppression of endogenous protein production should direct any isotopically labeled nutrients towards heterologous protein production, hopefully resulting in an isotopically labeled target protein within a natural abundance cellular background. By coordinating the inhibition of endogenous gene expression with exposure to isotopically labeled media and induction of heterologous protein expression, rifampicin could provide a means to specifically isotopically label a heterologously expressed protein.
Using membrane proteins involved in protein insertion (YidC) with a high level of recombinant expression and ion transport (KcsA) exhibiting low level of recombinant expression, we show that this approach can be used to target labels to proteins of interest, resulting in spectral simplification and decreased isotope labeling costs. Further, the method also can be adapted to suit different labeling schemes and protein expression levels, and can be used efficiently even when the amount of recombinant protein is significantly less than that of major endogenous MPs [which have copy numbers of ~105/cell (Renault et al. 2012b and references therein)]. By removing the need for high expression levels and purification of MPs, this method expands the applicability of ssNMR experiments to wider range of challenging MPs.
Cloning and recombinant protein expression
The gene for YidC from Escherichia coli (UniProt ID: P25714) was cloned into pHisLic vector with an N-terminal 6× histidine tag for expression under a T7 system using enzyme-free cloning, as described previously (de Jong et al. 2006). The gene for KcsA from Streptomyces lividans (UniProt ID: P0A334) was obtained in a pT7 vector, as described previously (van Dalen et al. 2000). E. coli LEMO21 cells (New England Biolabs) were transformed with these plasmids by heat shock before plating on LB with ampicillin (AMP) at 50 μg/mL and chloramphenicol (CAM) at 35 μg/mL. Cells were cultured according to standard practice (see also Baker et al. 2015). Precultures were grown in LB with AMP and CAM and then transferred to M9 minimal media (Folkers et al. 2004) supplemented with 1.0 g/L 14NH4Cl, 5.0 g/L 12C6-glucose, AMP and CAM overnight. Between 50 and 500 mL volumes of M9, supplemented as above, were inoculated with the overnight cultures to OD 0.1, and grown at 37 °C with shaking until reaching an OD ~1.5–2.0. Cells were harvested by centrifugation at 4000×g for 15 min before resuspension in equal volumes isotopically labeled supplemented M9 (1.0 g/L 15NH4Cl, 5.0 g/L 13C-glucose for uniformly labeled samples; 1.0 g/L 14NH4Cl, 5.0 g/L 12C-glucose and 200 mg/L each 15N and 13C labeled amino acid for specifically labeled samples). IPTG (isopropyl β-d-1-thiogalactopyranoside) at 0.5–1 mM final concentration was added to induce expression of the T7 polymerase and the cultures were incubated at 28 °C for 30 min with shaking. Rifampicin was added to a final concentration of 100 μg/mL and the cultures were incubated with shaking in the dark at 28 °C overnight. Cells were harvested by centrifugation at 4000×g for 15 min at 4 °C, and resuspended in 5–10 mL cold lysis buffer (20 mM Tris pH 7.4, 100 mM NaCl).
Cellular membrane isolation, inner membrane purification, and ssNMR sample preparation
Cells were lysed in a pressure cell homogenizer (Stansted) at 8000 psi without the addition of lysozyme. Cell debris was removed by centrifugation at 7000×g for 15 min, and any inclusion bodies were removed, where necessary, by centrifugation at 25,000×g for 15 min. Membranes were harvested by centrifugation at 100,000×g for 1 h.
Inner and outer membrane fractions were separated with sucrose gradients (Baker et al. 2015). Briefly, 2 mL of 55 % (w/v) sucrose, 8 mL of 51 % (w/v) sucrose, 8 mL of 45 % (w/v) sucrose, and 5 mL of 36 % (w/v) sucrose, all in 50 mM Tris pH 8.0 buffer, were poured in a stepwise manner, with 4 mL of sample mixed with sucrose to a final concentration of 20 % (w/v) layered on top. The gradients were centrifuged overnight in a SW32-Ti swinging bucket rotor (Beckman) at 100,000×g. Fractions corresponding to the outer membranes, inner membranes, and a mixture of outer and inner membranes were harvested with a syringe at the interface between the 55 and 51, 45 and 36, and 51 and 45 % sucrose layers, respectively. The band of mixed outer and inner membranes was observed after growth in M9 media, but not after growth in LB.
Membranes were washed twice with buffer (10 mM phosphate buffer pH 6.8 for YidC; 10 or 50 mM phosphate buffer pH 7.0, 10 or 50 mM potassium chloride, 10 or 50 mM sodium chloride, and 0.01 % sodium azide for KcsA inner and cellular membrane preparations, respectively) and collected by centrifugation at 125,000×g for 1– 2 h before packing into a 3.2 mm rotor for magic angle spinning (MAS).
Purification of reference proteoliposome samples
YidC with a hexa-histidine tag was over-expressed for purification as described for the cellular samples, with the following differences. After preculture, the main culture was inoculated in M9 with 13C6-glucose and 15NH4Cl at OD 0.1, and grown to OD 0.6 prior to induction with IPTG and 10 μM final concentration rhamnose. Rhamnose treated cells were found to produce more YidC when induced at lower ODs; no improvement was observed for induction at higher ODs like those used for rifampicin treatment. Cells were grown at 37 °C for a further 3–5 h until the OD was greater than 2.0. Cells were pelleted and lysed, and the membranes isolated as described for the cellular NMR samples. Membranes were solubilized at 4 °C by stirring with 2 % (w/v) dodecyl maltoside (DDM) (Anatrace) in 10 mM phosphate buffer (pH 6.8) containing 250 mM NaCl, 2 mM 2-mercaptoethanol, 10 % (v/v) glycerol, 0.03 % (w/v) DDM and 20 mM imidazole (Buffer A) for 2 h. Insoluble material was pelleted at 100,000×g for 1 h, and the supernatant was bound to 5 mL Ni–NTA resin (Qiagen) overnight. The resin was washed with 4 column volumes (CV) of buffer A, and then with 2 CV of buffer B (10 mM phosphate buffer pH 6.8, 100 mM NaCl, 2 mM 2-mercaptoethanol, 10 % (v/v) glycerol, 0.03 % (w/v) DDM and 20 mM imidazole). YidC was eluted with 5 CV of buffer B with 380 mM additional imidazole. The eluent was dialyzed four times for ~2 h against 75 mL of Buffer B without imidazole. The protein concentration was estimated using the BCA assay (Pierce) and then mixed with E. coli polar lipid extract (Avanti Polar Lipids), dissolved in water, at a ratio of 2 mg YidC: 1 mg lipid. DDM was removed by overnight incubation with BioBeads (BioRad) and YidC proteoliposomes were harvested by centrifugation at 100,000×g for 1 h. The proteoliposomes were washed in 10 mM phosphate buffer, pH 6.8, and pelleted at 125,000×g for ~2 h before being packed into a 3.2 mm MAS rotor.
Spectroscopy, processing, and referencing
All YidC samples were measured on a 700 MHz narrow-bore spectrometer (Bruker Biospin, Germany) with a 3.2 mm 1H, 13C, 15N MAS probe at 13 kHz MAS frequency, unless otherwise noted, and with a set temperature of 258 K (corresponding to an effective temperature of ~4 °C). Hartmann–Hahn 1H–15N and 1H–13C cross polarization (CP) used contact times of 650 and 500 μs, respectively, and 70–100 % ramps. 15N–13C SPECIFIC CP (Baldus et al. 1998) used a contact time of 4500 μs with optimized irradiation frequencies of 30.3 and 19 kHz, for 15N and 13C, respectively, and a ramp of 90–100 %. PARIS (Weingarth et al. 2009) was used for 13C–13C mixing, with a mixing time of 30 ms, 7.3 kHz 1H irradiation, and MAS frequency of 11 kHz. In all experiments, decoupling used SPINAL64 (Fung et al. 2000) with 1H irradiation of 83 kHz. 15N-edited 13C–13C detected experiments used Hartmann–Hahn 1H–15N CP followed by 15N–13Cα SPECIFIC CP and PARIS mixing.
KcsA samples were measured on an 800 MHz wide-bore spectrometer (Bruker BioSpin, Germany) with a 3.2 mm 1H, 13C, 15N MAS probe. Experiments were recorded at a set temperature of 253 K and MAS frequency of 12 kHz. CP contact times were 400 μs with a ramp of 70–100 % for 1H–15N, SPECIFIC CP used 3000 μs contact time with 90–100 % ramp, and irradiation of 57, 38, and 29 kHz was used for 1H, 15N, and 13C pulses, respectively. Decoupling used SPINAL64 with 83.3 kHz irradiation.
Spectra were referenced against adamantane (Harris et al. 2008) and histidine (Wei et al. 1999) powders. Data were processed with TopSpin 3.0 (Bruker Biospin) and analyzed using Sparky (Goddard and Kneller 2008). Chemical shift predictions were made with Shiftx2 (Neal et al. 2003) and FANDAS (Gradmann et al. 2012) using atomic model PDB ID 3wvf (Kumazaki et al. 2014).
Results and discussion
To remove signal from outer membrane proteins, we explored the use of inner membrane vesicles prepared from our cellular envelope samples. The sucrose gradient used to separate the inner and outer membrane resulted in a relatively low yield of inner membranes when cells were grown in M9 media, so ~200 mL of cell culture at OD ~2.0 was needed to fill a 3.2 mm MAS rotor. However, the spectral quality of 15N–13Cα SPECIFIC CP-based experiments using inner membrane samples of KcsA was considerably improved compared to using cellular membrane samples (Fig. 5b). We could readily observe resolved peaks with previous assignments (Schneider et al. 2008; van der Cruijsen et al. 2013) to the turret region (such as T61) and selectivity filter (T74, T75 or G79) of KcsA in the fully labeled inner membrane samples (Fig. 5b, c). Interestingly, we observed reduced ssNMR intensities for T74, for which we have recently found increased dynamics in the closed-conductive state (Koers et al. 2014). The same applies to G53 and A54, which exhibit stronger dynamics compared to other regions of the turret, such as T61 (van der Cruijsen et al. in preparation). Given that the lipid character influences the function of KcsA channels in lipid bilayers (Weingarth et al. 2013), these data provide initial evidence for structural conservation of KcsA irrespective of whether membrane mimetics such as asolectin that is rich in the zwitterionic phosphatidyl choline (PC) or E. coli inner membranes (rich in zwitterionic phosphatidyl ethanolamine, PE) are used.
Studying complex molecules in their biological context is an exciting prospect for structural biology. There is increasing evidence that the environment is an important contributor to the biophysical properties of proteins, modulating important biological processes such as ligand binding (e.g. Hubbard et al. 2003) and protein–protein interactions (e.g. Burz et al. 2006). In-cell NMR has even revealed different protein folding states in vitro and in vivo (Dedmon et al. 2002). For MPs, the impact of different environmental mimics has been well established in vitro. The difference between in vitro and in vivo experiments, however, is less well understood, due to the challenges of probing biological processes at an atomic level in a specific manner for these proteins. The work presented here expands the scope of cellular ssNMR to study the properties of MPs in their native environment. We have demonstrated that rifampicin treatment can be used to produce cellular membrane samples for ssNMR with specific protein labeling. Unlike in-cell NMR experiments where selective isotope incorporation was not found to significantly improve 15N experiments (Serber et al. 2001), we have demonstrated a clear improvement in ssNMR spectra of cellular membrane samples upon removal of background signals with rifampicin. This difference is most likely due to the use of direct 13C detection in our ssNMR experiments. These samples can be adapted to different isotopic labeling schemes and protein expression levels. As demonstrated with KcsA, individual resonances, important for function, can be observed. Although the rifampicin treatment can only be used to inhibit bacterial RNA polymerases (Campbell et al. 2001), a method similar to the single protein production (SPP) system (Suzuki et al. 2005) could potentially be used with eukaryotic cells.
Expression, purification and reconstitution of MPs into synthetic lipid bilayers can be a slow process, often requiring several weeks for more complicated samples. Purification necessitates removing MPs from their native environment, which potentially induces structural changes and loss of activity. Some membrane proteins are not amenable to established purification methods, and nearly impossible to purify in large quantities. As such, any method that allows MPs to be characterized in their native environment not only augments the biological relevance of any observations, but also presents an opportunity to study previously intractable systems. This system also naturally lends itself to the study of MP complexes, which are often disrupted by detergent solubilization.
From a spectroscopic point of view, rifampicin-treated cellular membrane samples present some unique advantages. Assignment of resonances in MPs can be challenging and dedicated labeling schemes are usually needed to reduce spectral overlap (see, for example, Etzkorn et al. 2007; Shi et al. 2011; Wang et al. 2013b; Sinnige et al. 2015). Specific labeling of a small number of amino acids presents an additional means to aid ssNMR assignment for these larger MPs. Due to the large number of samples that must be prepared to exhaustively cover the protein, this method would be extremely time and resource intensive for purified proteins. Here we have demonstrated that it is possible to study MPs isotopically labeled at a small number of amino acids from small cell culture volumes (~50 mL) without purification. This system does not rely on concentrating large cell cultures into small volumes before induction (as done in Mao et al. 2011), which in our hands results in lower expression levels of both membrane and soluble proteins. The rifampicin system presents a method by which to employ specific amino acid labeling quickly and inexpensively as an assignment strategy, potentially allowing ssNMR studies of membrane proteins which cannot be purified, of large membrane proteins with significant spectral overlap, and of membrane protein complexes.
We thank Prof. A. Killian for the pT7-KcsA plasmid. LAB was supported by a Human Frontier Science Program Long Term Fellowship. This work was supported by NWO (Grants 700.10.443 and 700.26.121 to M.B.).
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