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Cell-Free Protein Synthesis Enhancement from Real-Time NMR Metabolite Kinetics: Redirecting Energy Fluxes in Hybrid RRL Systems

  • Baptiste Panthu*
    Baptiste Panthu
    Univ. Lyon, ENS de Lyon, Univ. Claude Bernard, CNRS UMR 5239, INSERM U1210, Laboratory of Biology and Modelling of the Cell, 46 allée d’Italie Site Jacques Monod, F-69007 Lyon, France
    *E-mail: [email protected]
  • Théophile Ohlmann
    Théophile Ohlmann
    CIRI, Inserm, U1111, Université Claude Bernard Lyon 1, CNRS, UMR5308, ENS de Lyon, Univ. Lyon, F-69007 Lyon, France
  • Johan Perrier
    Johan Perrier
    Univ. Lyon, CNRS, Université Claude Bernard Lyon 1, ENS de Lyon, Institut des Sciences Analytiques, UMR 5280, 5 rue de la Doua, F-69100 Villeurbanne, France
  • Uwe Schlattner
    Uwe Schlattner
    Univ. Grenoble Alpes, Laboratory of Fundamental and Applied Bioenergetics (LBFA), 38058 Grenoble cedex, France
  • Pierre Jalinot
    Pierre Jalinot
    Univ. Lyon, ENS de Lyon, Univ. Claude Bernard, CNRS UMR 5239, INSERM U1210, Laboratory of Biology and Modelling of the Cell, 46 allée d’Italie Site Jacques Monod, F-69007 Lyon, France
  • Bénédicte Elena-Herrmann*
    Bénédicte Elena-Herrmann
    Univ. Lyon, CNRS, Université Claude Bernard Lyon 1, ENS de Lyon, Institut des Sciences Analytiques, UMR 5280, 5 rue de la Doua, F-69100 Villeurbanne, France
    *E-mail: [email protected]
  • , and 
  • Gilles J. P. Rautureau
    Gilles J. P. Rautureau
    Univ. Lyon, CNRS, Université Claude Bernard Lyon 1, ENS de Lyon, Institut des Sciences Analytiques, UMR 5280, 5 rue de la Doua, F-69100 Villeurbanne, France
Cite this: ACS Synth. Biol. 2018, 7, 1, 218–226
Publication Date (Web):September 15, 2017
https://doi.org/10.1021/acssynbio.7b00280

Copyright © 2017 American Chemical Society. This publication is licensed under these Terms of Use.

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Abstract

A counterintuitive cell-free protein synthesis (CFPS) strategy, based on reducing the ribosomal fraction in rabbit reticulocyte lysate (RRL), triggers the development of hybrid systems composed of RRL ribosome-free supernatant complemented with ribosomes from different mammalian cell-types. Hybrid RRL systems maintain translational properties of the original ribosome cell types, and deliver protein expression levels similar to RRL. Here, we show that persistent ribosome-associated metabolic activity consuming ATP is a major obstacle for maximal protein yield. We provide a detailed picture of hybrid CFPS systems energetic metabolism based on real-time nuclear magnetic resonance (NMR) investigation of metabolites kinetics. We demonstrate that protein synthesis capacity has an upper limit at native ribosome concentration and that lower amounts of the ribosomal fraction optimize energy fluxes toward protein translation, consequently increasing CFPS yield. These results provide a rationalized strategy for further mammalian CFPS developments and reveal the potential of real-time NMR metabolism phenotyping for optimization of cell-free protein expression systems.

Cell-free protein synthesis (CFPS) systems have become central tools for protein research or biotechnology. (1,2) For example, they can be used to produce not only therapeutic proteins such as antibodies, vaccine components or cytokines, (3−6) but also key pharmaceutical targets for drug design assays and structural biology, (7) or viral particles. (8) CFPS also constitutes excellent opportunities for direct sampling and screening of molecules in a close-to-physiological context, e.g., for numerous in vitro translational assays such as ribosome display techniques. (9) CFPS has been at the origin of ground breaking studies, such as elucidation of the genetic code, (10) and has been instrumental in the discovery of biological pathways involved in mRNA translation, stability, processing and regulation (11) or viral replication. (8) Yet, the development of CFPS systems to their full potential has been hampered by the complexity of their molecular machinery, which involves many cellular processes and regulatory circuits. (12) Only few active CFPS systems have been successfully developed so far, with lysates derived from E. coli, wheat germ, insect cells or rabbit reticulocytes being the most commonly used. (1)E. coli and wheat germ can produce large amounts of a given protein and have been optimized and routinely used in continuous CFPS protocols. However, such plant- or bacteria-based CFPS systems suffer from problems related to expression, folding and post-translational modifications of eukaryotic proteins. Thus, expression of many key protein targets in their active form is still a challenge. (1) On the other hand, mammalian CFPS systems are still believed to provide the most adequate machinery for bona fide protein production and downstream processing. Disappointingly, rabbit reticulocyte lysate (RRL), the main commercialized mammalian CFPS system (along with Chinese hamster ovary (CHO) and HeLa systems), is so far not optimal to deliver high protein production yield, (1) neither in its untreated form (uRRL) nor in its S7 nuclease treated RRL counterpart. (13)
We and others have recently proposed a novel in vitro CFPS system (14−17) based on rabbit reticulocyte ribosome-free lysate supernatant (ribFreeRRL) complemented with different ratios of purified ribosomal fractions, either from rabbit reticulocytes (reconstituted RRL or recRRL) or from diverse other cell types (hybrid RRL) (Figure 1a). Using this approach, protein expression became detectable through radiolabeling or luciferase reporter gene expression, (14,15,18,19) which allowed to decipher canonical translation mechanisms in the context of viral subversion (18) and tissue development. (19) This hybrid RRL approach allows a wide range of biological and technical observations as it maintains cellular properties such as specific mRNA translation regulation. Interestingly, hybrid RRL and recRRL initiate translation with the same rate than native RRL but are capable to maintain it for several hours, while translation rapidly levels off after 30mn in RRL, an effect that is not due to mRNA stability. (15)

Figure 1

Figure 1. CFPS energy consumption monitoring by real-time NMR. (a) RRL-derived CFPS reconstitution approach and nomenclature used in this article. Ribosomal fractions are easily isolated through ultracentrifugation under sucrose cushion. (b) Typical 1H NMR spectrum of a translating nativeRRL mix. Active CFPS reactions (200 μL) were monitored in real time inside the NMR spectrometer at 30 °C. (c) Evolution of metabolite concentrations quantified by real-time NMR during 2 h and normalized to their initial or final concentration. (d) Protein production yield based on luciferase activity measurement through a Renilla reporter gene expression. Each batch is supplemented by 27 nM of in vitro transcribed mRNA. Error bars represent the standard deviation of three experiments.

Here, we show that this persistence of translation in hybrid or reconstituted RRL systems is not directly due to metabolite supply, as both RRL and hybrid RRL are equally supplied with amino acids and tRNA. However, using real-time NMR, we identify persistent free energy-consuming metabolic activities associated with the ribosome fraction that monopolize most of the energy supply, independently of protein synthesis. Our results evidence that the energetic metabolism is the most crucial factor for efficient CFPS, and demonstrate that energy supply is essential to sustain long lasting translation. Using concentrations of ribosomal fractions lower than their native ratios significantly optimizes this issue.

Materials and Methods

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DNA Constructs and In Vitro Transcription

The β-globin and CrPV IRESes p0-renilla were previously described. (20) The pGEM-Renilla vector was used as template to generate polyadenylated transcripts by digestion with EcoRI. The 80 nt region containing the T7 promoter was deleted by PvuII/BamHI digestion and the resulting vector was used to insert the different 5′-UTRs. The human β-globin 5′-UTR with the authentic initiation codon was obtained by hybridizing two synthetic oligodeoxyribonucleotides (Eurogentec) and cloned into the double-digested vector (PvuII and BamHI restriction sites and T7 promoter were added with the oligos) generating the β-globin p0-renilla vector. The Cricket paralysis virus (CrPV) IRES was obtained by polymerase chain reaction (PCR) using cDNA prepared from infected flies. (21) PCR products was digested by PvuII and BamHI and cloned in the double-digested vector (T7 promoter and restriction sites were added by PCR) generating CrPV IRES p0-renilla vector.
RNAs were transcribed using the T7 RNA polymerase from templates linearized at the AflII site. Uncapped CrPV-Renilla mRNAs were obtained by using 1 μg of linear DNA template, 20 U of T7 RNA polymerase (Promega), 40 U of RNAsin (promega), 1.6 mM of each ribonucleotide triphosphate, 3 mM DTT in transcription buffer (40 mM Tris-HCl (pH 7.9), 6 mM MgCl2, 2 mM spermidine and 10 mM NaCl). For capped β-globin-Renilla mRNAs, the rGTP concentration was reduced to 0.32 mM and 1.28 mM of m7GpppG cap analogue (New England Biolabs) was added. The transcription reaction was carried out at 37 °C for 2h and mRNAs were precipitated with ammonium acetate at 2.5 M final concentration. The RNA pellet was resuspended in 30 μL RNase free water and RNA concentration was determined by reading the absorbance using Nanodrop technology. RNAs integrity was checked by electrophoresis on nondenaturing agarose gel.

Cell Culture

HeLa, 293T and MEF cells were obtained originally from American Tissue Type Culture Collection and were typically grown in DMEM containing 10% fetal calf serum supplemented with 50 U/mL of penicillin, 50 μg/mL of streptomycin in a humidified atmosphere containing 5% CO2 at 37 °C.

Preparation of RRL and Cell Lysate

Homemade cell lysate. All following steps were performed at 4 °C. 80% confluent cells from 20 10 cm diameter dishes were collected by centrifugation at 1000g for 5 min, rinsed 3 times with PBS, resuspended in an isovolume of hypotonic buffer (Hepes 10 mM, CH3CO2K 10 mM, (CH3CO2)2Mg, 1 mM and DTT 1 mM) and lysed using a Potter–Elvehjem homogenizer. The lysate was then centrifuged at 16 000g for 10 min and the supernatant was collected to yield the S10 supernatant extract. S10 protein total concentration was determined by the Bradford method and stored at 20 mg/mL concentration at −80 °C.
RRL and homemade cell lysate complementation. The method used has been described previously: (22) 1 mL of commercial untreated RRL (RRL) (Promega) or homemade cell lysate were supplemented with 25 μM hemin (Fluka), 25 μg creatine kinase (Sigma-Aldrich), 5 mg phosphocreatine (Fluka), 50 μg of bovine liver tRNAs (Sigma-Aldrich) and 2 mM of d-glucose (Sigma-Aldrich).
The in vitro translational assay was realized by adding 27 nM of in vitro transcribed mRNAs to a final volume of 200 μL (50% v/v RRL or homemade cell lysate) supplemented with 75 mM KCl, 0.75 mM MgCl2 and 20 μM amino acids mix. The translation reaction was realized in the NMR spectrometer at 30 °C. Unless otherwise stated in the text, the gloRenilla construct was used for every protein translation assays.

Protein Quantification

The reaction was stopped by addition of renilla lysis buffer (PJK). Renilla activity was measured using the renilla luciferase Assay System (PJK) in a Mithras (Berthold technologies) with 50 μL substrate injection and 10 s of signal integration program. Absolute quantification of in vitro translated protein yields was calibrated by 35S labeling for the commercial RRL luciferase. [35S]-methionine labeled radioactive protein was translated in the presence of 20 μM of amino acids mix minus methionine and 5 μCi of [35S]-methionine (PerkinElmer) for 30 mn before the reaction was stopped by addition of SDS loading buffer. Samples were resolved on a 12% SDS-PAGE, dried and subjected to autoradiography for 12 h by the use of Kodak Biomax films (Fisher Scientific) and the signal was quantified by using a Molecular Dynamics PhosphoImager FLA 5100 (Fuji). Protein concentration was determined relative to a serial dilution of [35S]-methionine stock.

RRL Fractionation and Reconstitution

The strategy and nomenclature are illustrated in Figure 1a. All steps were performed at 4 °C. After centrifugation of 1 mL of RRL for 2 h 15 min at 75 000 rpm in a TLA 100.3 rotor (Beckmann), 900 μL of ribosome free RRL was collected, frozen and stored at −80 °C. The extent of ribosome depletion from reticulocyte lysate was checked by translating 27 nM of in vitro transcribed capped and polyadenylated globin-renilla mRNA in the ribosome free RRL and validated when no luciferase activity could be detected. The ribosomal pellet was then rinsed three times in buffer R2 (Hepes 20 mM, NaCl 10 mM, KCl 25 mM, MgCl2 1.1 mM, β-mercaptoethanol 7 mM) and resuspended in 100 μL of R2 buffer to get a 10× final concentration. For ribosome salt wash experiments, the ribosomal pellet was washed using a 1 M sucrose cushion prepared with a R2 buffer including 25 mM to 1000 mM KCl, as stated in the text. After 2 h of centrifugation at 240 000g, the pellet containing the ribosomes was resuspended in R2 buffer.
The reconstituted lysate is then assembled by mixing 100 μL of ribosome free RRL with a fraction of scale from 0.02 to 1× ribosomal pellet. Typically, the standard reaction for NMR contains 100 μL of ribosome free RRL with 20 μg ribosomal pellet (annotated 0.1×) in a final volume of 200 μL. Upon reconstitution, the translation mixture is supplemented with 75 mM KCl, 0.75 mM MgCl2 and 20 μM amino acids mix.

NMR Acquisition

Sample preparation was as follows: 20 μL of D2O and 2 μL of 100 mM trimethylsilyl propionate (TMSP) were added to 200 μL of CFPS mix. Solutions were mixed thoroughly and 200 μL were transferred to 3 mm NMR tubes. All the solutions were strictly kept on ice until NMR acquisition. All NMR experiments were carried out on a Bruker 600 MHz NMR spectrometer equipped with a 5 mm TCI cryoprobe. The temperature was controlled at 30.0 °C throughout the experiments. Time series of standard 1H 1D NMR pulse sequence Carr–Purcell–Meiboom–Gill (CPMG) with water presaturation (Bruker pulse program cpmgpr1d) were acquired. CPMG experiments were preferred in this study to focus on small molecules by removing the important background contribution of large molecules such as proteins and nucleic acids. 32 free induction decays (FIDs) were collected with 48 074 data points and an acquisition time of 1.99 s. The relaxation delay was set to 4 s. The total acquisition time for each spectrum was 3 min 27 s. The acquisition was repeated during 2 to several hours. Identification of the metabolites was carried out from the 1D NMR data using the software ChenomX NMR Suite 7.0 (ChenomX Inc., Edmonton, Canada) and confirmed from analysis of 2D 1H–1H TOCSY, 1H–13C HSQC and J-Resolved NMR spectra recorded with standard parameters. The measured chemical shifts were compared to reference shifts of pure compounds using the HMDB (23) database. Metabolite concentrations were determined by manual fitting of the proton resonance lines for the compounds available in the ChenomX database. For comparison purposes, we normalized the data quantified by 1H NMR in CFPS reactions as fold changes using the following equation: X(t) = C(t)/Cref, in which Cref is respectively the initial or final concentration, whether the metabolite is consumed or produced during the reaction.

Results and Discussion

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Energetic Metabolism Evolution in RRL

In order to study metabolic activities and the related energy conversions occurring in translating RRL reactions, we used a quantitative real-time NMR strategy to monitor intact, translating samples over two to 6 h. As a primary energy source, an ATP regenerating system was used that is commonly applied for eukaryote CFPS, namely supplementation with 10 mM phosphocreatine (PCr) in the presence of creatine kinase (CK: ADP + PCr → ATP + Cr), with an additional 2 mM glucose supplementation. From the complex CFPS 1H NMR fingerprint (Figure 1b), a number of signals could be unambiguously assigned to specific metabolites and the evolution of their relative concentration followed over time, characterizing production, consumption, or steady-state behavior. This included PCr consumption and creatine production, but also the time course of other energy intermediates (ATP, GTP, glucose), metabolites and cellular constituents (e.g., Figure 1c,d).
Classical RRL translation (native RRL) was run as a control experiment. It revealed that the given PCr pool could maintain maximal energy supply for about 60 min, thus keeping ATP and also GTP concentrations globally stable during this time period (Figure 1c). At that time point, glucose was also entirely exhausted, and a drop in ATP and GTP occurred. Protein yield indicated very active translation during the first 30 min, which then slowed down and ceased after 45 min (Figure 1d). The GTP necessary for translation was apparently provided by an endogenous and nonlimiting nucleoside diphosphate kinase reaction (NDPK: GDP + ATP → GTP + ADP), as seen by the coupling of ATP and GTP concentrations (Figure 1c and Supporting Figure S1), a well documented process (24,25) that occurs in the cytoplasm. (26) All these kinetics are similar to those observed in prokaryotic systems, (27) except that we did not detect changes in α-ketoglutarate, consistent with the absence of mitochondria in our system, (28) and we observed production of pyruvate and lactate (data not shown). These data evidenced a considerable energy flux occurring during CFPS.

Energy Consumption Is Mostly Independent from Protein Production Yield of the Native RRL

To better understand energy expenditure during CFPS, we conducted experiments in which protein synthesis was blocked by cycloheximide (CHX), which inhibits the elongation process, and RNase A, to induce mRNA degradation. Surprisingly, at 60 min of reaction time (end of linear regime), PCr consumption was reduced by less than 20% (Figure 2a) as compared to the control RRL reaction. A slight reduction of glucose consumption was also tentatively detected, though the very low signal-to-noise at long reaction times for this metabolite prevents conclusion of a significant trend. This clearly demonstrated that translation was not the main consumer of ATP in the CFPS system. However, pyruvate and lactate production was reduced (data not shown), indicating that inhibition of translation reduces glycolytic flux.

Figure 2

Figure 2. Energy consumption in CFPS is maintained even when translation is blocked. (a) Phosphocreatine and glucose evolution during 2 h of in vitro translation: comparison of a nativeRRL control (△) versus an elongation-blocked nativeRRL (□) by addition of cycloheximide (CHX) and RNase A (top panel); comparison between a nativeRRL control (△), and without tRNA and amino acids (aa) supplementation (□) (middle panel); nativeRRL control (△) versus nativeRRL without glucose (□) or without CK and phosphocreatine (○) supplementation (bottom panel). (b) Protein production yield based on luciferase activity measurement through a Renilla reporter gene expression in nativeRRL (left) or without addition of amino acids and tRNA, glucose, CK, phosphocreatine or ribosomal fraction (ribosome). Absolute luminescence intensity is plotted on left y-axis. Concentration was determined using methionine 35S labeling and is plotted on right y-axis. Error bars represent the standard deviation of three experiments. (c) Comparison of a nativeRRL translating (△) versus a ribFreeRRL (□) system: phosphocreatine and glucose evolution during 2 h of in vitro translation. Error bars represent the standard deviation of three experiments.

Major Energy Consuming Processes Associate with the Ribosomal Fraction

To assess the origin of translation-independent energy consumption, we designed further experiments. First, we diminished regeneration of aminoacyl-tRNAs, which consumes at least 3 ATP per amino acid/tRNA coupling (Supporting Figure S2), by avoiding tRNA and amino acid additions to the reaction mixture. As compared to control RRL experiments, this decreased protein yield by about 20% (Figure 2b; in accordance with ref (29)). PCr consumption in the absence de tRNA/aa show slightly different behaviors for independent RRL batches, probably indicating variations of endogenous tRNA and amino acids concentrations in RRLs (Figure 2a; Supporting Figure S3). We observe either no difference or a maximum of 20% PCr consumption reduction in the absence de tRNA/aa supplementation (evaluated at 60 min reaction time), which indicates that, at most, 20% of the PCr available in a standard RRL reaction was used to regenerate aminoacyl-tRNAs. Altogether, at most 40% of the energy (a conservative overestimation) appeared to be spent for translation-related processes.
Next, we investigated the respective importance of PCr and glucose for protein yield. Removing PCr (or CK) led to an immediate drop in ATP and GTP levels (Supporting Figure S1), which precluded glucose utilization (Figure 2a; Supporting Figure S3) and protein synthesis (Figure 2b). In the context of high competition for ATP in RRL, and in the absence of an efficient ATP regenerating system, the low net ATP production of glycolysis is insufficient to fuel both glycolysis initial steps and competitive RRL metabolism. Consistently, glucose removal had no apparent effect on steady state ATP and GTP levels (Supporting Figure S1), confirming that the PCr/CK system is the primary and essential energy source, but interestingly glucose reduction lowered protein synthesis by about 30% (Figure 2b). Glycolysis thus appears to contribute to protein translation, either through NADH regeneration or via production of intermediate metabolites that may indirectly benefit translation. Notably, this is contrary to bacterial lysates, where glucose can be used as the major source of energy. (30,31) Finally, we conducted an experiment that omitted the ribosomal fraction, thus avoiding protein synthesis (Figure 2b). Under these conditions, PCr and glucose were still consumed, but at a much lower rate compared to the native RRL (Figure 2c) or to native RRL after CHX inhibition of protein synthesis (Figure 2a; Supporting Figure S3). Therefore, translation-independent free energy consumption seems to be largely driven by components copurified with the ribosomal fraction, or by the ribosome itself (Supporting Figure S1).
Our data suggest that energy fluxes during a CFPS reaction are complex. While a significant amount of free energy appears to be directed toward protein synthesis (at most 40%), a majority supports alternative pathways that are associated with the ribosomal fraction. This ribosomal fraction has recently been shown to be strikingly complex, with more than 1000 proteins associated with the ribosome machinery, constituting the riboproteome. (32) Those proteins are believed to create in vivo a favorable microenvironment necessary for driving, regulating, and securing specificity of the translation process. Interestingly, the riboproteome contains also many enzymes involved in common metabolic pathways, (32−34) consistent with the free energy-consuming processes that we found associated with the ribosomal fraction.

Riboproteome Dilution Is a Compromise between Translation-Independent Metabolism and Protein Synthesis

We explored the effect of riboproteome concentration on both translation-independent metabolism and protein synthesis in our CFPS system. Reducing the amount of the ribosomal fraction in ribosome free RRL (recRRL; Figure 1a) directly correlated with reduced free energy consumption of the system (Figure 3a), while protein synthesis was reduced in a more complex manner. In fact, reducing the ribosomal fraction of the original lysate (1×) to only 10% (0.1×) increased protein yield after 90 min of synthesis by about 4-fold (Figure 3b; Supporting Figure S4). Further reduction of the ribosomal fraction to 2% (0.02×) reduced protein yield. Under this condition, the yield was nonetheless higher than the one of the 1× controls. At native concentrations, the translation apparatus seems therefore saturated, an effect previously observed in vitro. (14,35) These differences in protein yield were not apparent at shorter reaction times (30 min), suggesting similar initial translation rates (Figure 3b; Supporting Figure S4). In fact, the different mixtures differed in the time course, with nativeRRL and 1× recRRL ceasing protein synthesis after 30–45 min, while recRRL, depending on the ribosome dilution, remained productive for longer reaction times.

Figure 3

Figure 3. Lowering the ribosome ratios increases the protein production yield. (a) Phosphocreatine and glucose evolution during 2 h of in vitro translation in nativeRRL (△), ribFreeRRL (□), recRRL with 0.02× (○), 0.1× (×), 0.5× (−) and 1× (+) ribosomal fraction addition. (b) Protein production yield evaluated as luciferase activity by luminometry at different time of incubation (30, 45, 60, and 90 min). Protein production was normalized to nativeRRL (RRL). RecRRL was supplemented with an increasing amount of ribosomal fraction from 0.02× to 1×. (c) recRRL comparison of protein production for 2 different constructs with ribosomes washed with 10 mM KCl to 1 M KCl through sucrose cushion. Error bars represent the standard deviation of three experiments. Absolute luminescence intensity is plotted on left y-axis. Concentration was determined using methionine 35S labeling and is plotted on right y-axis.

These results suggest that the metabolic activities associated with the ribosomal fraction contribute to the reduced translation yield in CFPS reactions by competing for energy supply. Using lower ribosomal fraction ratios than native systems appeared to be very beneficial: dilution of the competing free energy-consuming processes liberated energy reserves, which were then efficiently used by the ribosome translational machinery because of its saturated capacity down to concentration ratios of 0.1×. This conclusion is consistent with the longer-lasting ATP availability observed in the PURE system developed by Shimizu et al. (36) However, translation of this approach to eukaryotic CFPS seems not feasible, since individual components of the translation machinery have to be purified, which already limits large-scale commercial application of the E. coli system. (1)
We evaluated whether ribosome purification by high salt washes is advantageous as an alternative approach to deal with competing energy-consuming activities in the ribosomal fraction. Ribosome fractions were washed with up to 1 M KCl then rinsed with a 25 mM KCl buffer before being resuspended in assay buffer to avoid any effect of residual salt on translation efficiency (see Materials and Methods and ref (14)). More stringent washes during the ribosomes purification steps (using more than 350 mM, and up to 1 M KCl) progressively led to a loss of such ATP-consuming processes as compared to unwashed controls (Supporting Figure S5). Unfortunately, at the efficient KCl concentrations, the protein yield rapidly dropped by 50% at 350 mM KCl to even 90% at 500 mM KCl (Figure 3c) when using glo-Renilla mRNA (a capped and polyadenylated mRNA-dependent translation control). The result was slightly different when using the CrPV-Renilla construct (an mRNA that displays a viral intergenic IRES that does not need any eIF factors to be translated), with a maximal protein yield observed at 350 mM KCl, and a drop occurring only at higher KCl concentrations (Figure 3c). These experiments suggest that washes with low salt concentration can already dissociate not only metabolic enzymes, but also initiation factors and potentially other important translation components.

Evaluation of the Hybrid CFPS Approach Using Ribosomal Fractions from Various Cell Types

Finally, we studied such hybrid RRL approaches by using ribosomal fractions of different cell types (HeLa, MEF and HEK-293T cells) and comparing them to total cell lysates (Figure 1a). Protein yield was again quantified by Renilla protein production (Figure 4a). For experiments using HeLa and MEF cells, protein yield after 30 min was two logs lower in the standard cell lysates as compared to the hybrid RRL systems, while both were similar in HEK-293T experiments. For all three cell lines, protein yield after 60 min further increased in the hybrid systems, but not in the total cell lysates. As reported earlier, HeLa lysates display a very low protein production capacity (2 logs lesser than native RRL), (14) while the hybrid RRL complemented with a HeLa ribosomal fraction shows protein expression yields similar to native RRL for short incubation time (30 to 45 min) and similar to recRRL for longer incubation time. Similar results were observed for MEF cell lysate and hybrid RRL with MEF ribosomal fraction. On the opposite, both HEK-293T standard cell lysate and hybrid RRL performed well at short incubation time, demonstrating that the low yields obtained with HeLa and MEF lysates were not caused by our experimental conditions or chemicals. We further compared energy availability between cell lysates and the hybrid RRL with reduced ribosomal fraction ratio (equivalent to 0.1× RRL) similar as above. For HeLa lysates, glucose and PCr half-lives were only 5 min and 12 to 15 min respectively (Figure 4b). In striking contrast, the HeLa hybrid CFPS half-lives for glucose and PCr were increased to 55 and 85 min, respectively. This indicates that in our experimental setup the energy sources in the HeLa cell lysate (15 mM PCr, 2 mM glucose) are unable to fuel protein synthesis for much longer than 20–30 min, while in the hybrid CFPS they last for more than 2 h. Similar results were obtained for MEF and 293T cell lysates (Figure 4b). In comparison, HeLa cell lysates showed by far the fastest consumption of energy reserves, i.e., of ATP-consuming metabolic reactions, consistent with their cancer cell origin. We note that the duration of CFPS reactions is highly dependent on the energy supplies and experimental conditions. (37)

Figure 4

Figure 4. Application of the hybrid strategy to mammalian cell based CFPS delivers protein production levels comparable to RRL by energy consumption optimization. (a) Protein production yield after 30 and 60 min of in vitro translation at 30 °C for HeLa (purple, left), MEF (blue, middle), 293T (green, right) cell lysates and corresponding hybrid RRL. Error bars represent the standard deviation of three experiments. Absolute luminescence intensity is plotted on left y-axis. Concentration was determined using methionine 35S labeling and is plotted on right y-axis. (b) Phosphocreatine and glucose evolution determined by real-time NMR during 2 h of in vitro translation in cell lysates (△) and in corresponding hybrid RRL (×). HeLa were run in duplicate experiments, MEF and 293T as single experiments.

Altogether, we emphasize here the cell-specific energy consuming activities associated with the ribosomal fraction as crucial factors for protein translation yield. Dilution of the ribosomal fraction limits the waste of energy by translation-independent ATP-consuming pathways, promotes significantly longer lasting CFPS reactions, and thus maximizes ribosome activity and protein yield. While competing energy-consuming activities of the ribosomal fraction do not appear to contribute to translation, they may in fact be necessary for translation in vivo, but appear nonlimiting under the conditions tested. Manipulating the ribosome ratio was already used to enhance protein production in yeast CFPS, (35) but the underlying energetic aspects were not investigated. Alternative approaches to circumvent the energy issue would be to engineer the metabolic pathways or to specifically eliminate the futile cycles that are likely to drain energy resources in cellular lysates. While these strategies have proven successful for prokaryotes, (36,38) adapting them to eukaryotes appears so far not feasible due to the lack of information on the involved competing free energy consuming processes. The hybrid system approach investigated here appears therefore as an accessible, solid and pragmatic tool to improve protein yield. In fact, by emphasizing the importance of energy resource management, it could present a breakthrough for the elaboration of cell free protein synthesis systems. Both the hybrid approach and real-time NMR follow-up of CFPS appear as experimental strategies adaptable to a large variety of mammalian cell types, paving the way for the development of novel eukaryotic CFPS systems and versatile platforms for high yield protein production, riboproteome investigation, or gene expression studies.

Supporting Information

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The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acssynbio.7b00280.

  • Figures S1–S6 (PDF)

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Author Information

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  • Corresponding Authors
    • Baptiste Panthu - Univ. Lyon, ENS de Lyon, Univ. Claude Bernard, CNRS UMR 5239, INSERM U1210, Laboratory of Biology and Modelling of the Cell, 46 allée d’Italie Site Jacques Monod, F-69007 Lyon, FrancePresent Address: CIRI, Inserm, U1111, Université Claude Bernard Lyon 1, CNRS, UMR5308, ENS de Lyon, Univ. Lyon, F-69007, Lyon, France Email: [email protected]
    • Bénédicte Elena-Herrmann - Univ. Lyon, CNRS, Université Claude Bernard Lyon 1, ENS de Lyon, Institut des Sciences Analytiques, UMR 5280, 5 rue de la Doua, F-69100 Villeurbanne, FranceOrcidhttp://orcid.org/0000-0002-0230-1590 Email: [email protected]
  • Authors
    • Théophile Ohlmann - CIRI, Inserm, U1111, Université Claude Bernard Lyon 1, CNRS, UMR5308, ENS de Lyon, Univ. Lyon, F-69007 Lyon, France
    • Johan Perrier - Univ. Lyon, CNRS, Université Claude Bernard Lyon 1, ENS de Lyon, Institut des Sciences Analytiques, UMR 5280, 5 rue de la Doua, F-69100 Villeurbanne, France
    • Uwe Schlattner - Univ. Grenoble Alpes, Laboratory of Fundamental and Applied Bioenergetics (LBFA), 38058 Grenoble cedex, France
    • Pierre Jalinot - Univ. Lyon, ENS de Lyon, Univ. Claude Bernard, CNRS UMR 5239, INSERM U1210, Laboratory of Biology and Modelling of the Cell, 46 allée d’Italie Site Jacques Monod, F-69007 Lyon, France
    • Gilles J. P. Rautureau - Univ. Lyon, CNRS, Université Claude Bernard Lyon 1, ENS de Lyon, Institut des Sciences Analytiques, UMR 5280, 5 rue de la Doua, F-69100 Villeurbanne, FranceOrcidhttp://orcid.org/0000-0002-1064-0293
  • Notes
    The authors declare no competing financial interest.

Acknowledgments

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We are indebted to Dr. Neil Ferguson for long-term support, critical reading of the manuscript and stimulating discussions. The authors wish to thank the SFR Biosciences Lyon Gerland for contributing to the development of the project.

References

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This article references 38 other publications.

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  • Abstract

    Figure 1

    Figure 1. CFPS energy consumption monitoring by real-time NMR. (a) RRL-derived CFPS reconstitution approach and nomenclature used in this article. Ribosomal fractions are easily isolated through ultracentrifugation under sucrose cushion. (b) Typical 1H NMR spectrum of a translating nativeRRL mix. Active CFPS reactions (200 μL) were monitored in real time inside the NMR spectrometer at 30 °C. (c) Evolution of metabolite concentrations quantified by real-time NMR during 2 h and normalized to their initial or final concentration. (d) Protein production yield based on luciferase activity measurement through a Renilla reporter gene expression. Each batch is supplemented by 27 nM of in vitro transcribed mRNA. Error bars represent the standard deviation of three experiments.

    Figure 2

    Figure 2. Energy consumption in CFPS is maintained even when translation is blocked. (a) Phosphocreatine and glucose evolution during 2 h of in vitro translation: comparison of a nativeRRL control (△) versus an elongation-blocked nativeRRL (□) by addition of cycloheximide (CHX) and RNase A (top panel); comparison between a nativeRRL control (△), and without tRNA and amino acids (aa) supplementation (□) (middle panel); nativeRRL control (△) versus nativeRRL without glucose (□) or without CK and phosphocreatine (○) supplementation (bottom panel). (b) Protein production yield based on luciferase activity measurement through a Renilla reporter gene expression in nativeRRL (left) or without addition of amino acids and tRNA, glucose, CK, phosphocreatine or ribosomal fraction (ribosome). Absolute luminescence intensity is plotted on left y-axis. Concentration was determined using methionine 35S labeling and is plotted on right y-axis. Error bars represent the standard deviation of three experiments. (c) Comparison of a nativeRRL translating (△) versus a ribFreeRRL (□) system: phosphocreatine and glucose evolution during 2 h of in vitro translation. Error bars represent the standard deviation of three experiments.

    Figure 3

    Figure 3. Lowering the ribosome ratios increases the protein production yield. (a) Phosphocreatine and glucose evolution during 2 h of in vitro translation in nativeRRL (△), ribFreeRRL (□), recRRL with 0.02× (○), 0.1× (×), 0.5× (−) and 1× (+) ribosomal fraction addition. (b) Protein production yield evaluated as luciferase activity by luminometry at different time of incubation (30, 45, 60, and 90 min). Protein production was normalized to nativeRRL (RRL). RecRRL was supplemented with an increasing amount of ribosomal fraction from 0.02× to 1×. (c) recRRL comparison of protein production for 2 different constructs with ribosomes washed with 10 mM KCl to 1 M KCl through sucrose cushion. Error bars represent the standard deviation of three experiments. Absolute luminescence intensity is plotted on left y-axis. Concentration was determined using methionine 35S labeling and is plotted on right y-axis.

    Figure 4

    Figure 4. Application of the hybrid strategy to mammalian cell based CFPS delivers protein production levels comparable to RRL by energy consumption optimization. (a) Protein production yield after 30 and 60 min of in vitro translation at 30 °C for HeLa (purple, left), MEF (blue, middle), 293T (green, right) cell lysates and corresponding hybrid RRL. Error bars represent the standard deviation of three experiments. Absolute luminescence intensity is plotted on left y-axis. Concentration was determined using methionine 35S labeling and is plotted on right y-axis. (b) Phosphocreatine and glucose evolution determined by real-time NMR during 2 h of in vitro translation in cell lysates (△) and in corresponding hybrid RRL (×). HeLa were run in duplicate experiments, MEF and 293T as single experiments.

  • References

    ARTICLE SECTIONS
    Jump To

    This article references 38 other publications.

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    2. 2
      Schinn, S. M., Broadbent, A., Bradley, W. T., and Bundy, B. C. (2016) Protein synthesis directly from PCR: progress and applications of cell-free protein synthesis with linear DNA. New Biotechnol. 33, 480487,  DOI: 10.1016/j.nbt.2016.04.002
    3. 3
      Kanter, G., Yang, J., Voloshin, A., Levy, S., Swartz, J. R., and Levy, R. (2007) Cell-free production of scFv fusion proteins: an efficient approach for personalized lymphoma vaccines. Blood 109, 33933399,  DOI: 10.1182/blood-2006-07-030593
    4. 4
      Yang, J., Kanter, G., Voloshin, A., Levy, R., and Swartz, J. R. (2004) Expression of active murine granulocyte-macrophage colony-stimulating factor in an Escherichia coli cell-free system. Biotechnol. Prog. 20, 16891696,  DOI: 10.1021/bp034350b
    5. 5
      Yang, J., Kanter, G., Voloshin, A., Michel-Reydellet, N., Velkeen, H., Levy, R., and Swartz, J. R. (2005) Rapid expression of vaccine proteins for B-cell lymphoma in a cell-free system. Biotechnol. Bioeng. 89, 503511,  DOI: 10.1002/bit.20283
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      Zawada, J. F., Yin, G., Steiner, A. R., Yang, J., Naresh, A., Roy, S. M., Gold, D. S., Heinsohn, H. G., and Murray, C. J. (2011) Microscale to manufacturing scale-up of cell-free cytokine production--a new approach for shortening protein production development timelines. Biotechnol. Bioeng. 108, 15701578,  DOI: 10.1002/bit.23103
    7. 7
      Terada, T., Murata, T., Shirouzu, M., and Yokoyama, S. (2014) Cell-free expression of protein complexes for structural biology. Methods Mol. Biol. 1091, 151159,  DOI: 10.1007/978-1-62703-691-7_10
    8. 8
      Machida, K. and Imataka, H. (2015) Production methods for viral particles. Biotechnol. Lett. 37, 753760,  DOI: 10.1007/s10529-014-1741-9
    9. 9
      Pluckthun, A. (2012) Ribosome display: a perspective. Methods Mol. Biol. 805, 328,  DOI: 10.1007/978-1-61779-379-0_1
    10. 10
      Nirenberg, M. W. and Matthaei, J. H. (1961) The dependence of cell-free protein synthesis in E. coli upon naturally occurring or synthetic polyribonucleotides. Proc. Natl. Acad. Sci. U. S. A. 47, 15881602,  DOI: 10.1073/pnas.47.10.1588
    11. 11
      Mathonnet, G., Fabian, M. R., Svitkin, Y. V., Parsyan, A., Huck, L., Murata, T., Biffo, S., Merrick, W. C., Darzynkiewicz, E., Pillai, R. S., Filipowicz, W., Duchaine, T. F., and Sonenberg, N. (2007) MicroRNA inhibition of translation initiation in vitro by targeting the cap-binding complex eIF4F. Science 317, 17641767,  DOI: 10.1126/science.1146067
    12. 12
      Jackson, R. J., Hellen, C. U., and Pestova, T. V. (2010) The mechanism of eukaryotic translation initiation and principles of its regulation. Nat. Rev. Mol. Cell Biol. 11, 113127,  DOI: 10.1038/nrm2838
    13. 13
      Pelham, H. R. and Jackson, R. J. (1976) An efficient mRNA-dependent translation system from reticulocyte lysates. Eur. J. Biochem. 67, 247256,  DOI: 10.1111/j.1432-1033.1976.tb10656.x
    14. 14
      Panthu, B., Decimo, D., Balvay, L., and Ohlmann, T. (2015) In vitro translation in a hybrid cell free lysate with exogenous cellular ribosomes. Biochem. J. 467, 387398,  DOI: 10.1042/BJ20141498
    15. 15
      Panthu, B., Mure, F., Gruffat, H., and Ohlmann, T. (2015) In vitro translation of mRNAs that are in their native ribonucleoprotein complexes. Biochem. J. 472, 111119,  DOI: 10.1042/BJ20150772
    16. 16
      Penzo, M., Carnicelli, D., Montanaro, L., and Brigotti, M. (2016) A reconstituted cell-free assay for the evaluation of the intrinsic activity of purified human ribosomes. Nat. Protoc. 11, 13091325,  DOI: 10.1038/nprot.2016.072
    17. 17
      Penzo, M., Rocchi, L., Brugiere, S., Carnicelli, D., Onofrillo, C., Coute, Y., Brigotti, M., and Montanaro, L. (2015) Human ribosomes from cells with reduced dyskerin levels are intrinsically altered in translation. FASEB J. 29, 34723482,  DOI: 10.1096/fj.15-270991
    18. 18
      Panthu, B., Terrier, O., Carron, C., Traversier, A., Corbin, A., Balvay, L., Lina, B., Rosa-Calatrava, M., and Ohlmann, T. (2017) The NS1 protein from influenza virus stimulates translation initiation by enhancing ribosome recruitment to mRNAs. J. Mol. Biol.  DOI: 10.1016/j.jmb.2017.04.007
    19. 19
      Mazelin, L., Panthu, B., Nicot, A. S., Belotti, E., Tintignac, L., Teixeira, G., Zhang, Q., Risson, V., Baas, D., Delaune, E., Derumeaux, G., Taillandier, D., Ohlmann, T., Ovize, M., Gangloff, Y. G., and Schaeffer, L. (2016) mTOR inactivation in myocardium from infant mice rapidly leads to dilated cardiomyopathy due to translation defects and p53/JNK-mediated apoptosis. J. Mol. Cell. Cardiol. 97, 213225,  DOI: 10.1016/j.yjmcc.2016.04.011
    20. 20
      Soto-Rifo, R., Rubilar, P. S., Limousin, T., de Breyne, S., Decimo, D., and Ohlmann, T. (2012) DEAD-box protein DDX3 associates with eIF4F to promote translation of selected mRNAs. EMBO J. 31, 37453756,  DOI: 10.1038/emboj.2012.220
    21. 21
      Chtarbanova, S., Lamiable, O., Lee, K. Z., Galiana, D., Troxler, L., Meignin, C., Hetru, C., Hoffmann, J. A., Daeffler, L., and Imler, J. L. (2014) Drosophila C virus systemic infection leads to intestinal obstruction. J. Virol. 88, 1405714069,  DOI: 10.1128/JVI.02320-14
    22. 22
      Soto Rifo, R., Ricci, E. P., Decimo, D., Moncorge, O., and Ohlmann, T. (2007) Back to basics: the untreated rabbit reticulocyte lysate as a competitive system to recapitulate cap/poly(A) synergy and the selective advantage of IRES-driven translation. Nucleic Acids Res. 35, e121,  DOI: 10.1093/nar/gkm682
    23. 23
      Wishart, D. S., Jewison, T., Guo, A. C., Wilson, M., Knox, C., Liu, Y., Djoumbou, Y., Mandal, R., Aziat, F., Dong, E., Bouatra, S., Sinelnikov, I., Arndt, D., Xia, J., Liu, P., Yallou, F., Bjorndahl, T., Perez-Pineiro, R., Eisner, R., Allen, F., Neveu, V., Greiner, R., and Scalbert, A. (2013) HMDB 3.0--The Human Metabolome Database in 2013. Nucleic Acids Res. 41, D801807,  DOI: 10.1093/nar/gks1065
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