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A simple protocol for the production of highly deuterated proteins for biophysical studies

Highly deuterated protein samples expand the biophysics and biological tool kit by providing, among other qualities, contrast matching in neutron diffraction experiments and reduction of dipolar spin interactions from normally protonated proteins in magnetic resonance studies, impacting both electron paramagnetic resonance and NMR spectroscopy. In NMR applications, deuteration is often combined with other isotopic labeling patterns to expand the range of conventional NMR spectroscopy research in both solution and solid-state conditions. However, preparation of deuterated proteins is challenging. We present here a simple, effective, and user-friendly protocol to produce highly deuterated proteins in Escherichia coli cells. The protocol utilizes the common shaker flask growth method and the well-known pET system (which provides expression control via the T7 promotor) for large-scale recombinant protein expression. One liter expression typically yields 5 to 50 mg of highly deuterated protein. Our data demonstrate that the optimized procedure produces a comparable quantity of protein in deuterium (2H2O) oxide M9 medium compared with that in 1H2O M9 medium. The protocol will enable a broader utilization of deuterated proteins in a number of biophysical techniques.

Highly deuterated protein samples expand the biophysics and biological tool kit by providing, among other qualities, contrast matching in neutron diffraction experiments and reduction of dipolar spin interactions from normally protonated proteins in magnetic resonance studies, impacting both electron paramagnetic resonance and NMR spectroscopy. In NMR applications, deuteration is often combined with other isotopic labeling patterns to expand the range of conventional NMR spectroscopy research in both solution and solid-state conditions. However, preparation of deuterated proteins is challenging. We present here a simple, effective, and userfriendly protocol to produce highly deuterated proteins in Escherichia coli cells. The protocol utilizes the common shaker flask growth method and the well-known pET system (which provides expression control via the T7 promotor) for largescale recombinant protein expression. One liter expression typically yields 5 to 50 mg of highly deuterated protein. Our data demonstrate that the optimized procedure produces a comparable quantity of protein in deuterium ( 2 H 2 O) oxide M9 medium compared with that in 1 H 2 O M9 medium. The protocol will enable a broader utilization of deuterated proteins in a number of biophysical techniques.
Perdeuteration coupled with isotope labeling has been widely applied and greatly appreciated in the past 2 decades for protein NMR spectroscopy (1)(2)(3)(4)(5). Conventional NMR spectroscopy has benefited from uniformly labeled 15 N, 13 C protein samples, and, when these labeling patterns are combined with replacement of all nonexchangeable protons with deuterons to create a perdeuterated background, it is possible to study very large proteins and protein complexes (6)(7)(8)(9)(10). Furthermore, extension of the labeling pattern to include specific methyl group labeling ( 13 CH 3 ), within the context of an otherwise perdeuterated background, enables a still wider range of studies into the structures, binding interactions, and conformational dynamics of proteins and protein complexes (11)(12)(13). The combined effect of these sophisticated isotope labelings has permitted researchers tremendous breadth in gaining insights of biological systems and their dynamic interactions, using NMR spectroscopy as the main tool. In recent years, as the biophysical research toolbox expands rapidly, both in terms of technology and instrumentation, it has become very clear that deuteration of proteins and other biomolecular samples will greatly aid in high-quality outcomes for other biophysical applications, particularly small-angle neutron scattering (SANS) (14)(15)(16)(17), pulsed double echo electron resonance (DEER) techniques (18)(19)(20), and neutron reflectometry (21,22). The general requirement is to effectively remove, or make nonresponsive, a component of the measurement as a result of perdeuteration, either through contrast matching as in SANS (23) or elongation of relaxation times via removal of dipolar coupling as in DEER and NMR (3,7,24,25). The SANS and NMR methods are further enhanced in the context of segmental labeling (26)(27)(28)(29), wherein portions of a macromolecule may be prepared with deuteration for SANS (and/or isotopic labeling for NMR) and the other portions are protonated. Hence, the need for an efficient, reproducible protocol for high-level deuteration is broadly applicable to numerous experimental methods.
Although highly desired, deuterated ( 2 H) sample preparation has been notoriously problematic. Most researchers find it arduous and inconsistent. Bacterial cells (Escherichia coli) are the preferred host, as insect and mammalian cells are difficult to culture in a D 2 O medium (30)(31)(32). Even bacteria can be difficult to adapt to grow in high levels of D 2 O (33). Conventional procedures are time consuming and labor intensive, and the yield of deuterated samples, where the percentage labeling is >90%, can be very low. In our laboratory, we previously examined two commonly used methods: (1) medium switch prior to induction and (2) multistep media switch adaptation. In method (1), cells were grown in LB medium to the appropriate density for induction (A 600 0.8), pelleted by centrifugation, resuspended in 100% D 2 O medium, and induced immediately. This method often leads to complete cell death during induction, due to lack of proper adaptation. If cells do survive, the level of protein expression is very low. In method (2), the multistep media switch adaptation, cells were grown and transferred, stepwise, from LB to 50%, 75%, and 100% D 2 O media, which discards considerable isotopic media in the adaptation phase. This method also results in low protein expression due to multiple centrifugation and resuspension steps, rendering a generally unhealthy cell culture. The approximate yield of deuterated proteins from these methods is less than 25% of the yield from our current protocol reported here. When expression efficiency is low, the combined cost of D 2 O, 15 N/ 13 C sources, and/or specific 13 C-labeled metabolic precursor compounds can become very high, thus limiting widespread application and realization of the full potential.
We report a highly reproducible, user friendly, time and isotope efficient, and high percentage labeling deuteration procedure that can support the ever-expanding applications in the extended communities. Our protocol eliminates many of the common roadblocks: (1) the carryover of any rich medium (LB, Superbroth, etc.) into the final culture; (2) multiple media transfers during cell training, which involves repetitive cell pelleting, resulting in long recovery time for bacterial cells and unhealthy growth; (3) extended growth period that requires lengthy in-person monitoring; (4) an unhealthy cell growth curve that often leads to low expression and even cell death.

Results
The key to successful cell growth in deuterated medium ( 2 H-M9; for abbreviations and media descriptions see Experimental procedures) is maintaining cell density in the exponential phase (Fig. 1A). The protocol keeps A 600 at or slightly above 0.2 each time a culture is initiated or transferred or following a dilution. This approach ensures that the cells are the most healthy and viable to adapt into a higher D 2 O percentage environment. The protocol is described for production of 1 liter 2 H-M9 expression medium. LB and 1 H-M9 cultures do not require training (adaptation). A modified simple protocol for expression in LB and 1 H-M9 is described in the Supporting information and illustrated in Figure 1B.
A schematic flow chart for expression of deuterated proteins in 2 H-M9 is illustrated in Figure 2. In general, the flask (tube) used for cell growth is 4 to 10 times the volume of the culture, which ensures proper aeration during growth. Cultures are shaken in a temperature-regulated incubator at 250 rpm. Antibiotics are chosen based on the vectors in which the genes are constructed and added to all media for a given protein expression. Carbenicillin is preferred over ampicillin for stability, and all media should be stored at 4 C.  37 C for 30 min after supplying additives and then induced with 1 mM IPTG for 3 h. Cells are harvested at the end of day 3. This 37 C induction modification is necessary based on our knowledge and extensive testing on these proteins. We find that a significant number of mammalian proteins overexpressed in E. coli cells exhibit some level of toxicity, which can lead to reduced or completely abolished expression when induced at room temperature overnight. We recommend researchers test both induction conditions for their proteins of interest.

Day 4
1. Record A 600 of the final culture and collect a 20-μl sample for SDS-PAGE analysis. 2. Harvest cells by centrifugation (10 min, 3000g) and either proceed with protein-specific purification or store the cell pellet at −80 C.
As demonstration of the generality of this protocol, we provide data for the expression of six different mammalian proteins. The proteins span a wide range of protein types and sizes, comprising a moderate-size single-domain ubiquitinconjugating E2 enzyme (Ube2g2) (24,35,36), an isolated domain of a more complex multidomain protein (PH domain of ASAP1) (21), a multidomain signaling protein (PZA of ASAP1) (37), a posttranslationally modified (via myristoylation) protein (myr-Arf1) (38), a complex multidomain protein with internal intrinsically disordered regions (gp78C) (35,36,39), and the amphipathic membrane scaffolding protein (MSPΔH5) used to build nanodisc membrane mimetic particles (40,41). Each of these systems have been expressed with high levels of deuteration (≥96%), combined with either 15 N or complex 13 CH 3 -methyl labels. These proteins vary in size, expression level, solubility, etc., proving the flexibility and efficiency of the protocol.
The comparative performance for the six proteins is demonstrated by parallel expression in three different media: (1) LB, (2) 1 H-M9( 15 N), (3) 2 H-M9( 15 N) (Fig. 3). It is observed that expression levels, when following this protocol, are nearly equivalent for 1 H-M9( 15 N) and 2 H-M9( 15 N), and expression is only moderately reduced from LB. Each protein was purified according to the published procedures, and mass spectral data were acquired to enable determination of labeling efficiency (Table 1). In each case, the 15 N-labeling efficiency corresponds closely to the 99% 15 N enrichment of the nitrogen source, 15 NH 4 Cl. Furthermore, the labeling efficiency of 2 H, in the nonexchangeable positions, exceeds 96% for all six proteins. Each of these proteins can be prepared and purified in 5-to 50-mg/l quantities (data not shown), thus supporting a wide variety of structural and biophysical studies. In addition, we have observed that the expression level and the percentage of deuteration are not affected by other supplementary additives (such as a-keto acid precursors) in the medium, provided the carbon source ( 12 C 6 2 H 7 -glucose or 13 C 6 2 H 7 -glucose) is ≥97% deuterated (data not shown).

Discussion
The described procedure (Fig. 2) has been employed in our laboratory for the past decade. We confidently report that we can express the 30+ proteins from a wide range of difficult systems in highly deuterated form at similar or slightly reduced level compared with the expression in rich medium such as LB or protonated 1 H-M9 ( 15 N) medium. One of the keys to this protocol is proper attention to cell viability and avoidance of cellular and metabolic stress. Our approach deviates from conventional adaptation methods that encompass multiple steps of cell pelleting, media transfer, and resuspension. All these treatments incur additional stress and slow down, or endanger, cell growth. Unlike other published methods (42), this protocol is significantly simplified, wherein only two types of culture media (15 ml LB and 1 liter 2 H-M9) are needed for each 1 liter production. We achieve such simplicity by starting a small, healthy LB culture and expanding (by simple dilution) the LB culture with 2 H-M9 medium in a stepwise fashion, while maintaining the growth within the exponential phase through all media adjustments. Another advantage of the protocol is that there is no carryover of any initial LB/H 2 O media into the final expression culture, since we complete adaptation to 100% D2O at the end of day 2, which leads to the high percentage of deuteration (Table 1). In addition, the protocol is time efficient, as the total time from initial cell culture in day 2 to the harvest on the morning of day 4 is approximately 48 h. The effective use of time is achieved by two elements: (1) starting with a fresh, healthy culture in LB (H 2 O) medium and (2) adapting cells to a higher D 2 O medium by stepwise addition of 2 H-M9 medium, thus eliminating multiple cell-pelleting steps.
Another protein deuteration method (43,44) has encouraged high cell density growth combined with reduced culture volume. While this approach may be desirable for reducing the cost of isotope labels and D 2 O, the high cell density growth (A 600 = 6.0-10.0) may incur unforeseeable cellular and metabolic stress and may not be commonly applicable for a wide range of proteins. Our protocol emphasizes healthy cell growth and is mindful of minimizing the waste of D 2 O. Overall, only 45 ml of 2 H-M9 medium is discarded during cell adaptation per liter of growth production.
Interestingly, by observing the induced protein band intensity (Fig. 3), we find that, on average, most proteins express at a similar level across the three media. The minor changes presumably result from the characteristics of each protein. Contrary to conventional wisdom, not all proteins show reduced expression in M9 medium. Although some do show reduced expression, such as ASAP1-PH, ASAP-PZA, and myr-Arf1, others such as Ube2g2, gp78c, and MSPΔH5 express slightly better in M9 medium than in LB. One possible explanation is cell toxicity caused by low-level, basal expression in rich media (often called "leaky vectors"), while in M9 medium, basal expression is largely suppressed.
These flexible, combined labeling strategies further expand both NMR (21,26) and SANS applications (16,45). For NMR applications, in addition to performing 15 N-labeling in the presence of deuteration, we can perform the full variety of methyl-specific labeling (12,34,46) that has become popular in NMR studies. These labeling schemes include stereospecific, uniform side-chain labeling or suppression of specific residue types. For example, the 1 H-13 C methyl transverse relaxation optimized spectroscopy (Fig. S1A) and the 1 H-15 N transverse relaxation optimized spectroscopy (Fig. S1B) spectra of 2 H, 15 N, ILV proS 13 CH 3 -labeled ASAP1 PZA illustrate the high spectral quality, sensitivity, and resolution that are afforded by the labeling scheme when precursors for methyl labeling are included. ASAP1 PZA is a 44.5-kDa multidomain protein that binds to membrane-associated Arf1 (21,37,38), and NMR studies of these interactions at a nanodisc membrane surface require high levels of deuteration, as a protonated sample would result in highly overlapping, very broad, and weak signals. The protocol and labeling scheme have also been combined with segmental labeling (26), wherein only one domain of the multidomain PZA is deuterated or 15 N and 13 C labeled. We have utilized highly deuterated proteins to minimize background relaxation and enable more accurate measurement of long-range, intermolecular paramagnetic relaxation effects (Zhang et al., in preparation and (21)), facilitate solid-state 1 H-detected NMR (2,47), and DEER experiments utilizing nanodisc particles (Fig. S2, and (19)). For SANS applications, it can be advantageous to adjust the percentage of deuteration to provide optimal contrast matching in multicomponent systems (23), including protein:protein complexes (16) and membrane systems (14,15). Since our protocol provides for complete control of 2 H-labeling for nonexchangeable hydrogen atoms, it will be possible to finetune the level of deuteration. Adjustment of the final 2 H 2 O concentration and the ratio of 1 H 7 -glucose to 2 H 7 -glucose in the medium can achieve a specific level of deuteration, which can be readily monitored by mass spectrometry of the expressed protein.
The protocol has proven general and enables the use of deuteration to enhance NMR and other biophysical methodologies. The protocol should facilitate and expand the application of perdeuteration in a range of biophysical experiments and enable a broad range of systems to be examined in exquisite detail. Such studies will lead to better functional understanding and mechanistic detail in many biological systems.

Protein expression
Each of the six proteins were expressed in E. coli BL21 star (DE3) cells according to the protocol presented in Figure 2. Any adjustment of conditions followed published procedures for the individual protein, as noted in Results.

Mass spectrometry
Mass spectrometry data were acquired on an Agilent 6130 Quadrupole LC/MS System (Agilent Technologies, Inc) equipped with an electrospray source, operated in positive-ion mode. Separation was performed on a 300SB-C3 Poroshell column (2.1 mm × 75 mm; particle size 5 μm). The analytes were eluted at a flow rate of 1 ml/min with a 5 to 100% organic gradient over 5 min and holding the organic phase A for 1 min. Mobile phase A contained 5% acetic acid in water and mobile phase B was acetonitrile. Data acquisition, data analysis, and deconvolution of mass spectra were performed using Open Lab Chem Station Edition software (version C.01.05). Samples of purified proteins were typically 5 μl of a 5 μM solution.

Solutions and abbreviations
1. M9: minimal medium described by Neidhart et al. (48), where the nitrogen source is derived from NH 4 Cl and the carbon source is derived from glucose. Both the nitrogen and carbon source can be supplied with 15 N or 13 C. In addition, the carbon source can be supplied as 12 C 6 1 H 7glucose, 13   Mass spectrometry data and isotopic labeling efficiencies a Mass represents the replacement of nonexchangeable protons with 2 H. All exchangeable -NH, -NH 2 , -SH, -OH, and -COOH protons are calculated as 1 H, representing back exchange during purification and mass spectrometry analyses. The mass spectrometry analyses were performed on an electrospray LC-MS that ensures full protonation of all exchangeable sites. The calculations were performed with the software package Protein Sequence Analysis, developed by Ira Palmer, NIH. b Predicted mass computed using the amino acid sequence and the software package Protein Sequence Analysis, developed by Ira Palmer, NIH. 5. 2 H-M9( 15 N): M9 minimal medium wherein the solvent is