Abstract
Protein degradation mediated by the ubiquitin-proteasome pathway regulates signaling events in all eukaryotic cells, with implications in pathological conditions such as cancer and neurodegenerative diseases. Detection of protein degradation is an elementary need in basic and translational research. In vitro degradation assays, in particular, have been instrumental in the understanding of how cell proliferation and other fundamental cellular processes are regulated. These assays are direct, quantitative and highly informative but also laborious, typically relying on low-throughput polyacrylamide gel-electrophoresis followed by autoradiography or immunoblotting. We present protein degradation on chip (pDOC), a MITOMI-based integrated microfluidic device for discovery and analysis of ubiquitin-mediated proteolysis. The platform accommodates microchambers on which protein degradation is assayed quickly and simultaneously in physiologically relevant environments, using minute amount of reagents. Essentially, pDOC provides a multiplexed, sensitive and colorimetric alternative to the conventional degradation assays, with relevance to biomedical and translational research.
Introduction
Protein degradation by the ubiquitin-proteasome system is a central regulatory module through which the level of proteins in all eukaryotic cells remains balanced. Deviation from the desired amount of each protein at any given moment can be detrimental to the cell, leading to dysfunctional tissues and a wide range of illnesses in human, including cancer, cystic fibrosis, and neurodegenerative diseases [1] [2].
The core cascade underlying ubiquitination involves three enzymes: The E1 enzyme covalently binds and activates the ubiquitin molecule for transfer to an E2 conjugating enzyme. Then, the ubiquitin-conjugated E2 interacts with an E3 ubiquitin-ligase enzyme, which catalyzes the transfer of ubiquitin molecules from the E2 to the target protein or to a second ubiquitin molecule, typically via an isopeptide bond to a lysine residue. Finally, a target protein that is covalently bound to a chain of ubiquitin moieties can be recognized by the proteasome for degradation [1, 2]. Hundreds of different E3 enzymes underlie the enormous functional reach and specificity of the entire ubiquitination process. With respect to cell proliferation and cell cycle regulation, the ubiquitin ligases anaphase-promoting complex/cyclosome (APC/C) and Skp1-Cullin-F-box protein complex (SCF) are particularly important [3–5]. The substrate specificity of both complexes is dependent on coactivators: Cdc2O and Cdh1 for the APC/C and one of several F-box proteins for the SCF, e.g., Skp2 and β-TrCP [6–8]. Overall, orderly proteolysis mediated by cell cycle regulated E3 enzymes ensures unidirectional cell cycle in all eukaryotes. [6–10].
Protein degradation, however, cannot be automatically inferred from ubiquitination; while some forms of ubiquitin chains trigger proteolysis, monoubiquitination and other forms of polyubiquitination regulate signaling cascades via proteasome-independent pathways [11]. Furthermore, ubiquitination can be reversed by enzymes called deubiquitinases in a manner that can prevent proteolysis [12]. On the flip side, proteasomal degradation may not always be coupled to ubiquitination [13]. Thus, protein degradation must be determined directly.
Protein degradation assays in cell-free extracts, also known as ‘cell-free systems’, have been instrumental in cell biology research, enabling direct and quantitative analyses of ubiquitin-mediated proteolysis in physiologically relevant environments. In fact, much of the cell cycle principles were discovered by monitoring the degradation of cell cycle proteins in extracts from frog eggs or cycling human cells (see for example [14–20]). The extensive use of these ‘degradation assays’ in today’s modern era, is a testament to their efficacy (see for example [21–23]). Interestingly, conventional degradation assays have never truly benefited from modern technologies, and still typically rely on gel-electrophoresis, autoradiography and large amount of biological material.
Integrated microfluidics and pneumatic microvalves paved the way to protein chips in which the arrayed proteins are freshly expressed in a physiological environment that maintain proper protein folding and activity [24]. The target proteins are expressed either on chip or externally, and subsequently immobilized to microchambers via a designated surface chemistry. Then, a large panel of direct colorimetric assays can be performed over thousands of microchambers, using minute amounts of reagents. In recent years, we developed several microfluidic devices based on mechanically induced trapping of molecular interactions (MITOMI), with which we discovered and detected i) protein interactions with DNA, RNA, proteins and viruses; and ii) protein post-translational modification (PTM), specifically, phosphorylation, autophosphorylation and ubiquitination [24–30].
The combination of integrated microfluidics, protein arrays and cell-free systems from healthy or pathological sources holds great potential in biomedical research and diagnostics. In this study, we utilize the MITOMI platform for protein degradation analyses. The proof of concept is demonstrated using cell extracts with APC/C-specific activity. The method, named pDOC (protein degradation on chip) provides a fast, sensitive and costeffective alternative to the classic method by which proteasome-mediated proteolysis has been assayed in vitro almost unvaryingly for nearly half a century.
Materials and methods
Plasmids
pCS2-Flag-FA vector was generated by annealing Flag tag oligos and ligating final fragment into pCS2-FA vector using BamHI and FseI restriction sites. pCS2-Flag-FA-Securin-GFP w.t and Δ64 variant plasmids were generated by cloning w.t or Δ64 Securin-GFP [27] into pCS2-Flag-FA vector, using FseI (5’) and AscI (3’) flanked primers. pCS2-GemininΔ27-GFP was generated by deleting amino acids 1-27 from Geminin-GFP [27] using QuikChange® Lightning mutagenesis kit (Agilent, 210513). The plasmids pCS2-Flag-FA-Geminin-GFP w.t and Δ27 variant were generated by cloning Geminin-GFP into pCS2-Flag-FA vector sing FseI and AscI restriction sites. pCS2-Flag-FA-p27-GFP was generated by replacing Geminin open reading frame (ORF) with p27 ORF using FseI and AgeI restriction sites and pCS2-FA-p27 as a template [27]. Flag-p27-myc fragment was generated by a two-step assembly PCR using pCS2-Flag-FA-p27-GFP template, a first primer set containing a c-Flag tag (5’) and a Myc tag (3’), and a second primer set containing a T7 promoter (5’) and a T7 terminator sequence (3’). All GFP-tagged proteins caried enhanced variant of GFP (eGFP).
Cell culture maintenance
NDB cells are based on the HEK293 cell line. A detailed description of this cell system can be found in Ref [18]. NDB and HeLa S3 (ATCC; #CCL-2.2) cells were maintained in tissue culture dishes containing Dulbecco’s Modified Eagles Medium (DMEM) supplemented with 10% fetal bovine serum, 2 mM L-glutamine, and 1% Penicillin-Streptomycin solution (Biological Industries; #01-055-1A, #04-001-1A, #03-020-1B, #03-031-1B). Cells were maintained at 37°C in a humidified 5% CO2-containing atmosphere. HeLa S3 cells were either cultured on dishes or in 1-l glass spinner flasks in suspension (80 rpm). NDB cells were cultured in the presence of 5μg/ml Blasticidin (Life Technologies; #A11139-03) to maintain the pcDNA6/TR plasmid carrying the ORF for Tet repressor.
Cell synchronization
For late-mitosis synchronization, NDB cells were cultured in 150 mm/diameter dishes. After reaching a confluency of about 75%, the cells were treated with 1 μg/ml Tetracycline (Sigma-Aldrich; #87128) for 22 hr and harvest for extract preparation. For S-phase synchronization, HeLa S3 cells were cultured in suspension for 72 h up to a concentration of approximately 5×105 cells/ml. Cells were then supplemented with 2 mM Thymidine for 22 hr, washed with DMEM (twice, 5 min, 250×g) and released into prewarmed fresh media (37°C) for additional 9 hr. Cell culture was then supplemented again with 2 mM Thymidine for 19 h before harvest for extract preparation.
Preparation of cell extracts
HeLa S3 extracts: S-phase Synchronous HeLa S3 cells were washed with ice-cold 1× PBS and lysed in a swelling buffer (20 mM HEPES, pH 7.5, 2 mM MgCl2, 5 mM KCl, 1 mM Dithiothreitol [DTT], and protease inhibitor cocktail [Roche; #11836170001]) supplemented with energy-regenerating mixture, E-mix (1 mM ATP, 0.1 mM ethylene glycol-bis [β-aminoethyl ether]-N,N,N’,N’-tetra acetic acid [EGTA], 1 mM MgCl2, 7.5 mM creatine phosphate, 50 μg/ml creatine phosphokinase). Cells were incubated on ice for 30 min and homogenized by freeze–thawing cycles in liquid nitrogen and passed through a 21-G needle for 10 times. Extracts were cleared by subsequent centrifugation (17,000 × g; 10 and 40 min), and stored at −80°C. NDB mitotic extracts: Tet-induced NDB cells were collected from 20-24 150 mm dishes by gentle wash with ice-cold PBS. Extracts were prepared as described for HeLa S3. For more details see [18][31].
In vitro expression of target proteins
Target proteins were in-vitro expressed using rabbit reticulocyte lysate (TNT-coupled reticulocyte system; Promega; #L4600, #L4610) supplemented with either 35S-methionine/S-L-cysteine mix (PerkinElmer; #NEG772002MC) for radiography detection or with untagged Methionine (Promega #L118A) and Green Lysine (FluoroTect™ GreenLys, Promega #L5001).
Off-chip Degradation Assay
Degradation assays were performed in 20 μl cell extract supplemented with 1 μl of 20× energy regenerating mixture (see above), 1 μl of 10 mg/ml Ub solution (Boston Biochem; #U-100H), and 1μl radiolabeled in vitro translated protein of interest. For a negative control, reaction mixture was supplemented with proteasome inhibitor MG132 (20 μM; Boston Biochem; #I-130). Reaction mixtures were incubated at 28°C, and samples of 4-5 μl were collected in 15-20 min intervals. Off-chip detection: Time-point samples were mixed with 4× Laemmli Sample Buffer (BIO-RAD #1610747), denaturized (10 min, 95°C), and resolved by SDS-PAGE. Gels were soaked in a Methanol/Acetic acid (10/7.5%) fixative solution for 20 min, dried in vacuum and heat, and exposed to phosphor screen (Fuji) for 24-72 hr. In vitro translated proteins were visualized by autoradiography using Typhoon FLA 9500 Phosphorimager (GE Healthcare Life Sciences). Signal intensity (corrected for background signal) was measured by ImageJ software and was normalized to the signal at t0. All plots were created using Microsoft Excel software, version 16.20. Mean and SE values were calculated from three or four independent degradation assays. On-chip detection: Time-point samples were immediately frozen in liquid nitrogen. Before detection, samples were thawed on ice, flown through the chip for 3-5 min, and immobilized to protein chambers under the ‘button’ valve (see ‘Surface chemistry’ below). Next, the ‘button’ valves were closed, allowing unbound material to be washed by PBS. The level of target proteins (before and after degradation reactions) were determined by 488 nm-excitation and an 535/25 nm emission filter. Protein level could also be measured by immunofluorescence using fluorescently labeled antibodies (anti-Flag-Alexa 647, #15009; Cell Signaling, Danvers, MA, USA). These antibodies were flowed into the device and incubated with the immobilized proteins under the ‘button’ for 20 min at RT. Unbound antibodies were mechanically washed by PBS following the closing of the ‘button’ valve. Here, target protein levels were determined by 633 nm-excitation and an 692/40 nm emission filter.
Device fabrication
The microfluidic device is made of two layers of PDMS. The silicon wafers are written by photolithography (Heidelberg MLA 150). Then after, the soft lithography phase is induced using silicon elastomer polydimethylsiloxane (PDMS, SYLGARD 184, Dow Corning, USA) and its curing agent to fabricate the microfluidic devices. The microfluidic devices are consisting of two aligned PDMS layers, the flow and the control layers which are prepared using different ratios of PDMS and its curing agent; 5:1 and 20:1 for the control and flow layers, respectively. The control layer is degassed and baked for 30 min at 80°C. The flow layer is initially spin coated (Laurell, USA) at 2000 rpm for 60 sec and baked at 80°C for 30 min. Next, the flow and control layers are aligned using an automatic aligner machine (custom made) under a stereoscope and baked for 1.5h at 80°C for final bonding. The two-layer device is then peeled off from the wafer and bound to a cover slip glass via plasma treatment (air, 30%, 30 sec).
Surface chemistry
Biotinylated-BSA (1 μg/μl, Thermo) is flowed for 25 min through the device, allowing its binding to the epoxy surface. On top of the biotinylated-BSA, 0.5 μg/μl of Neutravidin (Pierce, Rockford, IL) is added (flow for 20 min). The ‘button’ valve is then closed, and biotinylated-PEG (1 μg/μl, (PG2-AMBN-5k, Nanocs Inc.) is flowed over for 20 min, passivating the flow layer, except for the buttons area. Following passivation, the ‘button’ valve is released and a flow of 0.2 μg/μl biotinylated anti-GFP antibodies (Abcam; #ab6658, Cambridge, United Kingdom) or 0.01 μg/μl biotinylated anti-Flag antibodies (Cell Signaling; #2908S Danvers, MA, USA) were applied. The antibodies bound to the exposed Neutravidin, specifically to the area under the ‘button’, creating an array of anti-GFP – or anti-Flag tag. PBS buffer was used for washing in between steps. In the case of p27 immobilization, surface chemistry was performed with 0.2 μg/ml donkey anti-mouse whole IgG antibodies (#715-065-150, Jackson Immuno research laboratories, Maryland, USA) followed by 20 min flow of 6.5 μg/ml anti p27 antibodies (Santa Cruz biotechnology, Heidelberg Germany; #1641 mouse).
On-chip degradation assay
Flag-Securin-GFP (w.t and Δ64 mut) and p27-GFP IVT products were flowed into the chip and immobilized on the surface under the ‘button’ at the protein chambers via its GFP tag, following by PBS buffer wash and scanned. Next, the ‘button’ valves were opened and the extract reaction mixtures were incubated with the protein chambers for 60 min (30°C). During the reaction, the level of the remining target protein was determined by GFP signal every 15 min. The decline in GFP signal correlated with degradation. After background signal subtraction, GFP signals were normalized to the signal at t0 (value of 1) or between 1 (max signal) and 0 (min signal).
Image and data analysis
LS reloaded microarray scanner, GenePix7.0 (Molecular Devices) and ImageJ image analysis software were used for analysis and presentation of the images. The signal measured around the button valve was considered as the background, since no immobilization of proteins was expected there. Yet, some background signal is always detected, which results from non-specific attachment of antibodies to the device surface. We subtracted the background signal around the buttons in a ring the size of 2R with 2-pixel spacing (see supplementary material in [27]).
Immunoblotting
Protein samples were mixed with x4 Laemmli buffer, denatured (10 min, 96°C), and resolved on freshly made 10% acrylamide gel using a Tris-glycine running buffer. Proteins were then electro-transferred onto a nitrocellulose membrane (Bio-Rad; #162-0115) using Trans-Blot Turbo transfer system (Bio-Rad). Ponceau S Solution (Sigma-Aldrich; #81462) was used to verify transfer quality. Membrane was washed (TBS), blocked (5% skimmed milk in TBST), and incubated (RT, 1 hr) with antibody solution (2.5% BSA and 0.05% sodium azide in PBS) before blotted with anti-Securin (Abcam; #AB3305) primary antibody (RT, 2 hrs). Anti-mouse Horseradish peroxidase-conjugated secondary antibody was purchased from Jackson ImmunoResearch (#115-035-003). ECL Signal was detected using EZ-ECL (Biological Industries; #20-500-171).
Results
pDOC is based on a MITOMI device, an integrated microfluidic chip originally developed to quantify protein-ligand interactions at equilibrium [24, 32]. The basic design was modified to contain an array of 32 by 32 microcompartments. Each compartment is separated into two chambers and controlled by three valves: a ‘neck’ valve that controls the diffusion (mixing) of material from chamber I into the ‘Protein chamber’, in which a specific target protein is trapped; a ‘sandwich’ valve that separates between cell unites; and the MITOMI ‘button’ valve, which traps interacting molecules beneath it, thus taking a snapshot of the interaction at equilibrium (Figure 1A). The pDOC chip design includes separation of the master control of the three valves into sections of the chip that can be activated and controlled independently. Compared to the control of each valve type for the entire chip, this separation not only improves valve response, but more importantly, it permits time-response assays on chip. Within each section, experiments are performed in all cell units in parallel, enabling high-throughput applications of the kind shown in our previous devices [24–26, 33].
Similar to the classic degradation assays, target proteins for pDOC analyses are in vitro translated (IVT) using rabbit reticulocyte lysate that allows correct folding and PTM. However, quantitative detection is based on fluorescence rather than radioactivity. The IVT products are immobilized on the glass surface of the chip via biotin-avidin binding. To this end, specific biotinylated antibodies are applied under the button valve. Following pull down of the target protein the unbound material, such as reticulocyte lysate and cell extract, is washed away. Then the entire chip is passivated by PEG-Biotin, except for the area beneath the button (Figure. 1A; for more details, see our previous publications [25, 26, 33]. The freedom to control flow in individual sections enables multiple regimes of surface chemistry on one chip.
The device is compatible with multiple strategies of surface chemistry and possible experimental setups (illustrated in Figure 1B): i) The target protein is tagged on both N’ and C’ termini. One tag is used for immobilization via tag-specific biotinylated antibodies and the second tag is green fluorescent protein (GFP), which is used for detection. ii) The target protein is single-tagged with GFP, which is used for both immobilization and detection. iii) The target protein is double tagged. Here, however, detection is based on fluorescently labeled antibodies against short non-fluorescent tags (e.g., Flag). iv) The target protein is in vitro translated in lysate containing fluorescent lysine and immobilized by protein-specific antibodies. Importantly, immobilization can be performed with non-biotinylated antibodies if the surface chemistry also includes biotinylated IgG. Overall, the flexibility of the method simplifies assay optimization according to specific needs and limitations.
pDOC facilitates analysis of protein degradation in cell-free extracts
Conceptually, analysis of protein degradation by pDOC is direct, simple and fast; signal detection is based on in situ quantification of fluorescent signals, thus obviating gel-electrophoresis and any other gel-related procedures, e.g fixation, drying, autoradiography or immunoblotting, and long exposures. pDOC functions like a chromatography column; it isolates and concentrates the target protein on the protein chamber of each cell unit. Our first goal was to examine whether the signal sensitivity and dynamic range of pDOC enables time-based quantification of IVT products following incubation with cell extracts in tube. As a proof of concept, we utilized mitotic extracts from HEK293 cells that are blocked in an anaphase-like state due to high levels of non-degradable Cyclin-B1. This mitotic cell-free system, hereafter referred to as NDB, recapitulates APC/CCdc20-mediated proteolysis of the cell cycle proteins Securin and Geminin [18, 31].
Conventional degradation assays of radiolabeled Flag-Securin-GFP and Flag-Geminin-GFP (IVT products) in NDB mitotic extracts are shown in Figure 2A. Control experiments with non-degradable mutant variants (Geminin Δ27 and Securin Δ64) demonstrate the specificity of the assay. Equivalent experiments were performed with non-radioactive IVT products. First, we performed an end-point assay. After 60 min incubation, reaction samples were loaded on pDOC through separate channels and scanned for GFP fluorescence. Control reactions, in which IVT products were incubated in PBS, allowed us to normalize the level of each target protein at t60 min (extracts/PBS ratio) and to estimate background signals. Overall, on-chip detection demonstrates a sharp reduction in the level of Geminin and Securin following incubation in NDB mitotic extracts, whereas non-degradable variants remained stable, exhibiting ~80% of the control GFP signals in PBS. At this juncture, we noted that background signals from reticulocyte lysate, cell extracts, and non-specific immobilization were minor (Figure S1).
Next, we tested whether pDOC can be utilized to obtain reliable kinetic information on protein degradation. Flag-Securin-GFP and Flag-Geminin-GFP were incubated in NDB mitotic extracts for 60 min, and reaction samples were snap frozen in liquid nitrogen every 15 min. After quick thawing, samples representing five time-points were loaded on the chip for signal quantification. Comparable analyses were performed using SDS-PAGE and autoradiography. Non-degradable Flag-SecurinΔ64-GFP and Flag-GemininΔ27-GFP variants were also assayed by autoradiography for control. Considering the vast differences between the two methods of detections, GFP and autoradiography signals were normalized between 0 and 1, meaning that the signal at t60 min was subtracted from all other time points. The resulting values were normalized to max signal at t0 and plotted. As shown in Figure 1C and D, on-chip analysis by pDOC and off-chip analysis by SDS-PAGE-autoradiography exhibited near identical degradation patterns of Flag-Securin-GFP and Flag-Geminin-GFP. Note that reaction cocktails contained 1 μl IVT, 20 μl extracts and 2 μl ubiquitin/E. mix solution, following our standard protocol [18, 27, 34] For each time point, 5 μl reaction mixes were flowed for a period of 3 min through protein chambers with open MITOMI button valves, allowing immobilization of the target protein, while the neck valves were closed. This loading protocol enabled clear visualization of the target protein without the concern of signal saturation (Figure S2). We concluded that pDOC facilitates both end-point and time-course analyses of protein degradation in vitro.
Importantly, Flag-Securin-GFP was immobilized to protein chambers via biotinylated anti-GFP antibodies rather than anti-Flag antibodies. By doing so, we effectively demonstrated that GFP can serve for both immobilization and detection, eliminating the need for two tags. Overall, we find GFP to be an optimal tag for signal detection on-chip. The fusion of a fluorescent protein to a target protein, however, may distort protein folding in ways that effectively limit ubiquitination and proteolysis. In this context, the flexibility of pDOC is particularly advantageous. Figure 3A depicts three configurations by which Securin degradation was analyzed on chip. Securin level was measured at t0 and t60 min. Signal detection can be direct, either by GFP or green-Lys. While the former is brighter, the latter allows quantification without tagging. Indirect detection by immunofluorescence was found to be equally informative. Here, short immunodetectable tags minimize the risk of misfolding, but the target proteins must be double tagged because a single tag cannot be used for both immunolabeling and immobilization. Detection by immunofluorescence requires an additional 30 min but we benefit from the bright signal of the fluorophores to which plenty of commercial antibodies are coupled. All three detection protocols revealed the degradation of Securin in NDB mitotic extracts.
The versatility of pDOC was further demonstrated using tag-free p27 (Figure 3B). Degradation of p27 is mediated by SCFSkp2 E3 ligase, rather than APC/C, and is orchestrated with the DNA synthesis (S) phase of the cell cycle [35]. p27 degradation was assayed in extracts from S-phase synchronous HeLa S3 cells and analyzed by pDOC. The protein was immobilized via anti-p27 and biotinylated anti-IgG antibodies, and detected by green-Lys. Analysis by pDOC revealed the stereotypical instability of p27 in S-phase extracts (Figure 3B). The degradation pattern resembled that measured by autoradiography (Figure S3). Thus, pDOC can be utilized for degradation assays of untagged proteins. Furthermore, the method is specific and not restricted to a certain type of cell-free systems.
The advantages of autoradiography-based degradation assays include minimal signal-to-noise ratio, high specificity of the signal and linearity of the assay. However, this does not come without a cost. First, the 35S isotope is a short-lived reagent with half-life of ~3 months.
Second, on the gel the signal is spread over a well of 4-5 mm width (standard 10-well gel). Third, in a standard degradation assay, 1-2 μl IVT product is diluted in 20-30 μl cell extracts whose protein concentration is about 20-25 mg/ml. Thus, the amount of IVT loaded per lane is limited by the maximum separation capacity of the gel. De facto, we load 4-5 μl of reaction mix into a well of a standard 10-well mini-gel of 1 mm thick. Fourth, in vitro translation is challenging for large proteins because of ribosome processivity. Thus, although large proteins incorporate more radiolabeled Methionine/Cysteine relative to small proteins, the overall signal of the full-length protein can be impractical for reliable quantification. Fifth, the exposure time for a high-quality signal is typically within a range of 12-24 hrs. Note that the abovementioned points 2-4 are equally relevant when protein degradation is assayed by SDS-PAGE and immunoblotting. As for point 5, western blotting does not require long exposures. Yet, the overall incubation time with antibodies is long. Furthermore, IVT products in immunoblot-based assays must be tagged in order to be distinguished from the endogenous proteins in the extracts. At this juncture, it is important to note that whether degradation is assayed by autoradiography or immunoblotting, informative expression of IVT products must be validated beforehand, in itself, a day long procedure. Equivalent validation by pDOC is instantaneous.
On-chip, the fluorescence signals of Flag-Securin-GFP at t60 min were above background level and more noticeable compared to autoradiography (Figures 2A, 3A and S1). Yet, when the signal at t60 min was subtracted from all other time-points, the overall degradation patterns of Flag-Securin-GFP, as revealed by pDOC and autoradiography, was similar (Figure 2C and D). This observation suggests that pDOC detects protein residue still lingering in the reaction mix after 60 min incubation, and which are barely detected by autoradiography, if at all. Thus, it can be argued that the sensitivity of pDOC surpasses that of the conventional autoradiography, and if so, pDOC not only facilitates in vitro degradation assays, but also significantly reduces reagent consumption and cost per assay. To test that, we diluted Flag-Securin-GFP in reticulocyte lysate 4- and 10-fold and incubated the substrate in NDB mitotic extracts for 1 hr, while maintaining the original reticulocyte lysate /extract volume ratio of 1/20 (μl). Time point samples were analyzed by pDOC (based on GFP fluorescence). Equivalent experiments performed in parallel with 35S-labeled Flag-Securin-GFP, following the conventional assay. To clarify, radiolabeled and non-radiolabeled substrates were expressed simultaneously from the same TNT®/DNA solution mix. Furthermore, degradation assays performed a day after delivery of the 35S-Met/35S-Cys solution to our lab (~1175Ci/mmol), and the gels were exposed to phosphor screen overnight. Yet, whenever the IVT substrate was diluted, the signal obtained by autoradiography was below any acceptable standard, even at t0, and decreased to barely or undetectable levels after 15 min incubation (Figure 4A). Conversely, analyses by pDOC were informative in all three conditions (Figure 4B). We could detect bona fide signals of 4- and 10-fold diluted Flag-Securin-GFP at t0 as well as at t60 min, recording the full dynamics of the protein in NDB mitotic extracts. On a more practical note, we effectively demonstrated that protein degradation can be analyzed with 0.1 μl IVT product and 2 μl cell extracts, thereby saving 90% of the reagents. This feature is particularly valuable in assays which rely on limited biological material, e.g., extracts from primary cells, and normal/pathological tissue samples.
pDOC unveils remanent amount of Flag-Securin-GFP that could not be visualized by conventional methods. Interestingly, while the overall degradation pattern of Flag-Securin-GFP in all three conditions was similar, we noticed a systematic delay in Flag-Securin-GFP degradation as the substrate concentration was reduced. This observation has not been identified previously in our lab, and could be attributed to rate-limiting steps along the process of ubiquitination/degradation of Securin, and perhaps APC/C substrates overall [36, 37].Note that the estimated concentration of Flag-Securin-GFP in the reaction mix was 17 nM (before further dilution in reticulocyte lysate; Figure S4).
On-chip assay for protein degradation
Until now, we have demonstrated the capacity of pDOC to facilitate analysis of in tube protein degradation assays, i.e. the assay takes place prior to its introduction to the chip (Figures 2–4). However, our protocol enables a complete on-chip assay for protein degradation. Reaction samples are loaded on the chip while neck-valves are closed, i.e., without access to chamber I. Target proteins are then immobilized and captured at the protein chamber and remaining materials are washed away (Figure 1A). Technically, the neck valve allows trapping of a target protein in the protein chamber under the MITOMI button valve, and cell-free extracts in chamber I, noted here as ‘extract chamber’ (Figure 5A). The opening of the neck valve enables diffusion of cell extracts into the protein chamber. The motivation for an on-chip assay is threefold: 1) higher-throughput, especially if the target proteins are expressed on-chip, which is possible for MITOMI-based devices; 2) reagent-saving and cost per assay. In fact, 5 μl cell extracts are sufficient to fill a thousand cell units; 3) analysis of protein degradation in real time. The challenge, however, is the limited degradation capacity of <1 nl extracts per cell unit, which never been tested on any platform.
We decided to test the feasibility of protein degradation on chip. To this end, wt and non-degradable variants of Flag-Securin-GFP as well as Flag-p27-GFP (IVT products) were loaded on the chip and captured in protein chambers along separate channels. All proteins were immobilized via biotinylated anti-GFP antibodies. After washing, NDB mitotic extracts were loaded into the extract chamber and trapped by closing the neck valve. Untrapped materials were washed away. The device was heated to 30°C and scanned to obtain signals of t0. The opening of the neck valve initiated degradation reactions in all cell units simultaneously (Figure 5A and B), and the chip was scanned in time intervals of 20 min. While the fluorescent signal of non-degradable Securin and p27 remained stable in NDB mitotic extracts throughout the experiment, the signal of Flag-Securin-GFP diminished with time (Figure 5C), revealing the regulated proteolysis of this protein in anaphase.
Discussion
pDOC is a lab-on-chip platform devised to facilitate and simplify discovery and analysis of protein degradation in physiologically relevant contexts. The chip accommodates hundreds of microchambers in which protein degradation can be assayed promptly and simultaneously using considerably lower quantities of reagents compared to conventional assays. The latter feature is especially crucial for the cell-free extracts whose production can be a bottleneck in terms of time, activity, and amount. Signal detection on pDOC is based on in situ quantification of fluorescent signal. The method is independent of contaminating radioactive materials and yet, with sensitivity that surpasses traditional assays. A comparison between pDOC and the conventional degradation assay is illustrated in Figure 6.
The flexibility of pDOC is significant; the platform facilitates almost all possible experimental designs. First, both tagged and untagged proteins can be assayed, with detection based on incorporation of Green-Lys, fluorescent proteins, or immunodetectable tags. Second, pDOC can be used as an integrated microfluidics column for instant analyses of off-chip degradation reaction. Alternatively, protein degradation can be assayed entirely on-chip and in real time. Third, pDOC allows high-throughput experiments, in which the same protein is tested for degradation in multiple extract/reaction solutions, or the same extracts are applied to dozens of different proteins, simultaneously. Finally and importantly, pDOC is inherently compatible with on-chip in vitro translation [24, 25, 27]. By expressing array of proteins on-chip, one can multiplex the protein targets from tens to thousands, increasing throughput dramatically. The downside of on-chip expression is that the assembly of the device with the DNA microarray is not trivial and currently has to be performed by experts in specialized laboratories.
Ubiquitin-mediated proteolysis is routinely assayed in hundreds of research laboratories worldwide. We devised pDOC to facilitate and simplify in vitro analyses of protein degradation. The method is fast, sensitive, reagent-saving, cost-effective, and inherently optimal for both low- and high-throughput studies. It is also noteworthy that full automation of the platform is foreseeable. We therefore believe that pDOC holds a great potential in basic and translational research.
The authors declare no competing interests
Supplementary information
Acknowledgements
We thank the Gerber and Tzur lab members for sharing reagents. The Tzur lab is supported by the Israel Science Foundation (ISF) Grant no. 2038/19.
Footnotes
↵3 These authors jointly supervised this work