We thank the Natural Sciences and Engineering Research Council (NSERC) and the Canadian Cancer Society for financial support. SCCS thanks NSERC and VNL thanks the Ontario Graduate Scholarship (OGS) program for graduate fellowships. ARW thanks the CRC for a Canada Research Chair.

Part 1: Device Fabrication
<ol>
<li>Clean glass substrates in Piranha solution (3:1 conc. sulfuric acid:30% hydrogen peroxide). Leave the substrates in Piranha solution for 10 min with frequent agitation.</li>
<li>After rinsing in deionized (DI) water and drying the substrates with N2 gas, place the substrates inside the electron beam chamber for chromium deposition (thickness of 250 nm).</li>
<li>To dehydrate the chromium-coated substrate, rinse in isopropanol and then bake on a hot plate for 5 min at 115&deg;C.</li>
<li>Dry the substrates and prime with hexamethyldisilazane (HMDS) by spin-coating (30 s, 3000 rpm). Spin-coat again (using identical parameters) with Shipley S1811 photoresist.</li>
<li>Pre-bake the substrate on a hot-plate (100&deg;C, 2 min), then pattern the photoresist by exposure to ultraviolet (UV) irradiation for 5 s through a photomask.</li>
<li>Develop the UV-exposed substrates in Shipley MF 321 developer for 3 min and wash in DI water. Post-bake on a hot-plate at 100&deg;C for 1 min.</li>
<li>Etch the exposed chromium by immersing in chromium etchant for 30 s. Rinse in DI water, then immerse in AZ300T stripper for 10 min to remove the remaining photoresist. Rinse in DI water and dry with N2 gas.</li>
<li>Deposit 2-5 &mu;m Parylene-C (an insulating polymer) by chemical vapor deposition onto a substrate bearing patterned chromium. Deposit 50 nm of Teflon-AF (to make the surface hydrophobic) by spin-coating a solution (1% wt/wt in Fluorinert FC-40) at 2000 rpm for 60 s. Post-bake on a hot-plate (160 &deg;C, 10 min). </li>
<li>To form the top plate, coat an un-patterned indium tin oxide (ITO) glass substrates 50 nm Teflon-AF, as above.</li>
<li>Post-bake both substrates on a hot-plate (160 &deg;C, 10 min).</li>
</ol>
Part 2: Device Set-up and Automation
<ol>
<li>In digital microfluidic devices operated in "two-plate" configuration,1 droplets are positioned between a bottom substrate (with patterned electrodes) and a top substrate (with one contiguous, transparent electrode, typically formed from ITO).</li>
<li>To set the device up, remove polymer coatings from the contact pads of a bottom substrate by gentle scraping with a scalpel. Couple the exposed pads of the bottom substrate with 40-pin connectors (Figure 1a).</li>
<li>Power-on a computer running LabView (National Instruments, Austin, TX), a home-built control box (featuring an array of high-voltage relays that are controlled by signals from a National Instruments DaqPad), a function generator, and an amplifier. The computer/control box facilitates user control over application of 100 VRMS/18 kHz signals to the device via the 40-pin connectors.</li>
<li>Assemble the device by positioning two pieces of double-sided tape (140 &mu;m total thickness) on the edges of the bottom substrate and complete with unpatterned ITO slides (top plate).</li>
<li>To initialize the control system, run the LabView calibration program (written in-house) to calibrate the feedback control.</li>
<li>Load the sample and reagents into the appropriate reservoirs (see section 3, below) and enclose them under the top substrate (Teflon-coated side facing down). Attach the ground connector to the top substrate.</li>
<li>Load the actuation code in LabView (written in-house) and execute the program to initiate droplet actuation.</li>
</ol>
Part 3: Sample and Reagent Preparation
<ol>
<li>Prepare working buffer (WB): 100 mM TrisHCl (pH 7.8) and 0.08% Pluronic F127 w/v.</li>
<li>Dissolve protein(s) to be analyzed in WB and pipette sample into reservoir 1 (R-1) on the device, as shown in Figure 1b.</li>
<li>Prepare precipitant, 20% Trichloroacetic acid (TCA) in DI water, and pipette into R-2.</li>
<li>Prepare wash buffer, 70/30 Chloroform/acetonitrile (ACN) and pipette into R-4.</li>
<li>Prepare resolubilizing buffer, 100 mM Ammonium Bicarbonate (pH 8.0) and 0.08% Pluronic F127 w/v, and pipette into R-5.</li>
<li>Dissolve reductant, tris(2-carboxyethyl)phosphine (TCEP), at 10 mM in WB and pipette into R-6.</li>
<li>Dissolve alkylating agent, iodoacetamide (IAM), at 12 mM in WB and pipette into R-7.</li>
<li>Dissolve trypsin in WB at a concentration equal to1/5 of the concentration of the total protein sample (see 3.1, above) and pipette into R-8.</li>
</ol>
Part 4: Digital Microfluidic Sample Processing
<ol>
<li>The following procedure is implemented automatically by means of LabView code written in-house. The volume of droplets dispensed from reservoirs is defined by the dimensions of the electrodes (in this case, dispensed droplets are ~600nL).</li>
<li>Dispense a droplet of protein-containing sample from R-1 and a droplet of precipitant from R-2 (Figure 2, Frame 1). Merge the two droplets and allow the combined droplet to incubate for 5 min, resulting in the precipitation of proteins onto the surface (Figure 2, Frames 2-3). Actuate the supernatant away from the precipitated protein to the waste reservoir, R-3 (Figure 2, Frame 4).</li>
<li>Dispense three droplets of wash buffer from R-4 and drive them across the precipitated protein to the waste reservoir, R-3 (Figure 2, Frame 5).</li>
<li>Allow the precipitate to dry, and dispense a droplet of resolubilizing buffer from R-5 to the protein. Allow the buffer to incubate for 20 min until the precipitate has dissolved (Figure 2, Frame 6).</li>
<li>Dispense a droplet of reductant from R-6 and merge it with the sample droplet. Mix the combined droplet by actuating it across 6 electrodes in a circular pattern. Allow the droplet to incubate for 1 h at room temperature in a humidified chamber (Figure 3, Frames 1-2).</li>
<li>Dispense a droplet of alkylating agent from R-7 and merge it with the sample droplet, followed by mixing (as in 4.5). Allow the droplet to incubate for 15 min at room temperature in a humidified chamber, protected from light (Figure 3, Frames 2-3).</li>
<li>Dispense a droplet of trypsin from R-8, and merge it with the sample droplet, followed by mixing (as in 4.5). Allow the droplet to incubate for 3 h at 37&deg;C in a humidified chamber on a hot plate (Figure 3, Frames 3-4).</li>
</ol>
Part 5: Post-Processing Sample Preparation
<ol>
<li>Remove top substrate and quench digestion reaction by adding 0.6 mL of 2.5% trifluoroacetic acid in water.</li>
<li>Purify quenched reaction product using C18 ZipTips (Millipore, Billerica, MA) according to manufacturer&#x2019;s instructions.</li>
<li>Dilute purified sample in DI water to a final volume of 100 mL.</li>
</ol>
Part 6: Mass Spectrometry
<ol>
<li>Evaluate the processed sample on a nano-HPLC mated to a tandem mass spectrometer. The HPLC system used here is from Eksigent Technologies and is equipped with a trap column (3 mm dia. C18 beads, 150 &mu;m ID x 2.5 cm) and a reversed-phase separation column (3 &mu;m dia. C18 beads, 75 &mu;m ID x 15 cm). The mass spectrometer used here is an LTQ from Thermo-Fisher Scientific.</li>
<li>Load a 2 &mu;L sample onto the trap column at 5&mu;L/min in a mobile phase comprising 5% acetonitrile in water. Separate the sample on the reversed phase column at 0.5&mu;L/min using a linear gradient from 20% acetonitrile to 80% acetonitrile over 25 min. Continue running at 80% acetonitrile for another 10 min.</li>
<li>Analyze the bands eluting off the column by tandem mass spectrometry. In the work reported here, the eluent is sampled into the mass spectrometer via a nanoelectrospray emitter (20 &mu;m ID tapered to 10 &mu;m ID) from NewObjective.</li>
<li>Identify the proteins in the sample using a database search program such as SEQUEST. In this work, we used the version of SEQUEST incorporated in Bioworks v3.01 (Thermo-Fisher Scientific), and identified proteins had probability scores of less than 1.0E-03 (equivalent to 99.9% confidence interval), and sequence coverages of at least 30% (Figure 4).</li>
</ol>
<img src="/files/ftp_upload/1603/1603fig1a.jpg" align="Figure 1a" /> <img src="/files/ftp_upload/1603/1603fig1b.jpg" align="Figure 1b" /> Figure 1. (a) A picture a DMF device mated to 40-pin connectors for automated droplet actuation. (b) A schematic of a device depicting the positioning of sample and reagents required for a proteomic workup.
<img src="/files/ftp_upload/1603/1603fig2.jpg" align="Figure 2" /> Figure 2. Frames from a movie depicting the automated extraction and purification of BSA in 20% TCA (precipitant) and 70/30% chloroform/acetonitrile (rinse solution). In frame 6, the precipitated protein is redissolved in a droplet of 100 mM ammonium bicarbonate.
<img src="/files/ftp_upload/1603/1603fig3.jpg" align="Figure 3" /> Figure 3. Frames from a movie illustrating sequential reduction, alkylation, and digestion of a droplet of resolubilized protein. In this figure, the reagents are colored with dyes for clarity; in practice, the reagents are not colored.
<img src="/files/ftp_upload/1603/1603fig4.jpg" align="Figure 4" /> Figure 4. MS chromatogram of a sample of bovine serum albumin processed by digital microfluidics. 25 distinct peptides were identified (99.9% confidence interval) corresponding a sequence coverage of 44%.

<ol>
	<li>Abdelgawad, M., Wheeler, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Digital+Revolution:+A+New+Paradigm+for+Microfluidics.">The Digital Revolution: A New Paradigm for Microfluidics.</a> Adv. Mat. 21, 920-925 (2009).</li><li>Jebrail, M., Wheeler, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Digital+Microfluidic+Method+for+Protein+Extraction+by+Precipitation.">A Digital Microfluidic Method for Protein Extraction by Precipitation.</a> Anal. Chem. 81, 330-335 (2009).</li><li>Luk, V. N., Wheeler, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Digital+Microfluidic+Approach+to+Proteomic+Sample+Processing.">A Digital Microfluidic Approach to Proteomic Sample Processing.</a> Anal. Chem. 81, 4524-4530 (2009).</li></ol>

The lack of standardized sample handling and processing in proteomics is a major limitation for the field. In addition, conventional macroscale sample handling involves multiple containers and solution transfers, which can lead to sample loss and contamination. A potential solution to these problems is to form integrated systems for sample processing relying on digital microfluidics1 (DMF). In previous work, DMF was shown to be useful for efficient removal of unwanted contaminants in heterogeneous protein-cont...

digital microfluidics for automated proteomic processing

Clinical proteomics has emerged as an important new discipline, promising the discovery of biomarkers that will be useful for early diagnosis and prognosis of disease. While clinical proteomic methods vary widely, a common characteristic is the need for (i) extraction of proteins from extremely heterogeneous fluids (i.e. serum, whole blood, etc.) and (ii) extensive biochemical processing prior to analysis. Here, we report a new digital microfluidics (DMF) based method integrating several processing steps used in clinical proteomics. This includes protein extraction, resolubilization, reduction, alkylation and enzymatic digestion. Digital microfluidics is a microscale fluid-handling technique in which nanoliter-microliter sized droplets are manipulated on an open surface. Droplets are positioned on top of an array of electrodes that are coated by a dielectric layer - when an electrical potential is applied to the droplet, charges accumulate on either side of the dielectric. The charges serve as electrostatic handles that can be used to control droplet position, and by biasing a sequence of electrodes in series, droplets can be made to dispense, move, merge, mix, and split on the surface. Therefore, DMF is a natural fit for carrying rapid, sequential, multistep, miniaturized automated biochemical assays. This represents a significant advance over conventional methods (relying on manual pipetting or robots), and has the potential to be a useful new tool in clinical proteomics.
Mais J. Jebrail, Vivienne N. Luk, and Steve C. C. Shih contributed equally to this work.
Sergio L. S. Freire's current address is at the University of the Sciences in Philadelphia located at 600 South 43rd Street, Philadelphia, PA 19104.

Watch this Scientific Journal Video about Digital Microfluidics for Automated Proteomic Processing at JoVE.com

Digital Microfluidics for Automated Proteomic Processing

Digital Microfluidics is a technique characterized by the manipulation of discrete droplets (~nL - mL) on an array of electrodes by the application of electrical fields. It is well-suited for carrying out rapid, sequential, miniaturized automated biochemical assays. Here, we report a platform capable of automating several proteomic processing steps.

Clinical proteomics has emerged as an important new discipline, promising the discovery of biomarkers that will be useful for early diagnosis and ...

digital-microfluidics-for-automated-proteomic-processing

Research

JoVE Journal

Biology

12.4K Views.  University of Toronto. Digital Microfluidics is a technique characterized by the manipulation of discrete droplets (~nL - mL) on an array of electrodes by the application of electrical fields. It is well-suited for carrying out rapid, sequential, miniaturized automated biochemical assays. Here, we report a platform capable of automating several proteomic processing steps.

Video: Digital Microfluidics for Automated Proteomic Processing

Chemotactic Response of Marine Micro-Organisms to Micro-Scale Nutrient Layers

The degree to which planktonic microbes can exploit microscale resource patches will have considerable implications for oceanic trophodynamics and biogeochemical flux. However, to take advantage of nutrient patches in the ocean, swimming microbes must overcome the influences of physical forces including molecular diffusion and turbulent shear, which will limit the availability of patches and the ability of bacteria to locate them. Until recently, methodological limitations have precluded direct examinations of microbial behaviour within patchy habitats and realistic small-scale flow conditions. Hence, much of our current knowledge regarding microbial behaviour in the ocean has been procured from theoretical predictions. To obtain new information on microbial foraging behaviour in the ocean we have applied soft lithographic fabrication techniques to develop 2 microfluidic devices, which we have used to create (i) microscale nutrient patches with dimensions and diffusive characteristics relevant to oceanic processes and (ii) microscale vortices, with shear rates corresponding to those expected in the ocean. These microfluidic devices have permitted a first direct examination of microbial swimming and chemotactic behaviour within a heterogeneous and dynamic seascape. The combined use of epifluorescence and phase contrast microscopy allow direct examinations of the physical dimensions and diffusive characteristics of nutrient patches, while observing the population-level aggregative response, in addition to the swimming behaviour of individual microbes. These experiments have revealed that some species of phytoplankton, heterotrophic bacteria and phagotrophic protists are adept at locating and exploiting diffusing microscale resource patches within very short time frames. We have also shown that up to moderate shear rates, marine bacteria are able to  fight the flow  and swim through their environment at their own accord. However, beyond a threshold high shear level, bacteria are aligned in the shear flow and are less capable of swimming without disturbance from the flow. Microfluidics represents a novel and inexpensive approach for studying aquatic microbial ecology, and due to its suitability for accurately creating realistic flow fields and substrate gradients at the microscale, is ideally applicable to examinations of microbial behaviour at the smallest scales of interaction.  We therefore suggest that microfluidics represents a valuable tool for obtaining a better understanding of the ecology of microorganisms in the ocean.

The fabrication of microfluidic channels and their implementation in experiments for studying the chemotactic foraging behaviour of marine microbes within a patchy nutrient seascape and the swimming behaviour of bacteria within shear flow are described.

The degree to which planktonic microbes can exploit microscale resource patches will have considerable implications for oceanic trophodynamics and ...

Studies of Bacterial Chemotaxis Using Microfluidics - Interview

Applying Microfluidics to Electrophysiology

Microfluidics can be integrated with standard electrophysiology techniques to allow new experimental modalities.  Specifically, the motivation for the microfluidic brain slice device is discussed including how the device docks to standard perfusion chambers and the technique of passive pumping which is used to deliver boluses of neuromodulators to the brain slice.  By simplifying the device design, we are able to achieve a practical solution to the current unmet electrophysiology need of applying multiple neuromodulators across multiple regions of the brain slice.  This is achieved by substituting the standard coverglass substrate of the perfusion chamber with a thin microfluidic device bonded to the coverglass substrate.  This was then attached to the perfusion chamber and small holes connect the open-well of the perfusion chamber to the microfluidic channels buried within the microfluidic substrate.  These microfluidic channels are interfaced with ports drilled into the edge of the perfusion chamber to access and deliver stimulants.  This project represents how the field of microfluidics is transitioning away from proof-of concept device demonstrations and into practical solutions for unmet experimental and clinical needs.

Microfluidics can be integrated with standard electrophysiology techniques to allow new experimental modalities.  Specifically, the motivation for the ...

Development of New Therapeutic Applications Using Microfluidics

Electrophoretic Separation of Proteins

Electrophoresis is used to separate complex mixtures of proteins (e.g., from cells, subcellular fractions, column fractions, or immunoprecipitates), to investigate subunit compositions, and to verify homogeneity of protein samples. It can also serve to purify proteins for use in further applications. In polyacrylamide gel electrophoresis, proteins migrate in response to an electrical field through pores in a polyacrylamide gel matrix; pore size decreases with increasing acrylamide concentration. The combination of pore size and protein charge, size, and shape determines the migration rate of the protein. In this unit, the standard Laemmli method is described for discontinuous gel electrophoresis under denaturing conditions, i.e., in the presence of sodium dodecyl sulfate (SDS). 

In this video, we demonstrate a method for electrophoretic separation of proteins using poly-acrylimide gel electrophoresis (PAGE).

Electrophoresis is used to separate complex mixtures of proteins (e.g., from cells, subcellular fractions, column fractions, or immunoprecipitates), to ...

Immunoblot Analysis

Immunoblotting (western blotting) is a rapid and sensitive assay for the detection and characterization of proteins that works by exploiting the specificity inherent in antigen-antibody recognition. It involves the solubilization and electrophoretic separation of proteins, glycoproteins, or lipopolysaccharides by gel electrophoresis, followed by quantitative transfer and irreversible binding to nitrocellulose, PVDF, or nylon. The immunoblotting technique has been useful in identifying specific antigens recognized by polyclonal or monoclonal antibodies and is highly sensitive (1 ng of antigen can be detected). This unit provides protocols for protein separation, blotting proteins onto membranes, immunoprobing, and visualization using chromogenic or chemiluminescent substrates.

Immunoblotting (western blotting) is a rapid and sensitive assay for the detection and characterization of proteins that works by exploiting the specificity inherent in antigen-antibody recognition. This video provides protocols for protein separation, blotting proteins onto membranes, immunoprobing, and visualization using chromogenic or chemiluminescent substrates.

Immunoblotting (western blotting) is a rapid and sensitive assay for the detection and characterization of proteins that works by exploiting the ...

Staining Proteins in Gels

Following separation by electrophoretic methods, proteins in a gel can be detected by several staining methods. This unit describes protocols for detecting proteins by four popular methods. Coomassie blue staining is an easy and rapid method. Silver staining, while more time consuming, is considerably more sensitive and can thus be used to detect smaller amounts of protein. Fluorescent staining is a popular alternative to traditional staining procedures, mainly because it is more sensitive than Coomassie staining, and is often as sensitive as silver staining. Staining of proteins with SYPRO Orange and SYPRO Ruby are also demonstrated here.

Following separation by electrophoretic methods, proteins in a gel can be detected by several staining methods. Staining of proteins with Coomassie Blue, Silver Staining, SYPRO Orange, SYPRO Ruby are demonstrated in this video.

Following separation by electrophoretic methods, proteins in a gel can be detected by several staining methods. This unit describes protocols for ...

Toxin Induction and Protein Extraction from Fusarium spp. Cultures for Proteomic Studies

Fusaria are filamentous fungi able to produce different toxins. Fusarium mycotoxins such as deoxynivalenol, nivalenol, T2, zearelenone, fusaric acid, moniliformin, etc... have adverse effects on both human and animal health and some are considered as pathogenicity factors. Proteomic studies showed to be effective for deciphering toxin production mechanisms (Taylor et al., 2008) as well as for identifying potential pathogenic factors (Paper et al., 2007, Houterman et al., 2007) in Fusaria. It becomes therefore fundamental to establish reliable methods for comparing between proteomic studies in order to rely on true differences found in protein expression among experiments, strains and laboratories. The procedure that will be described should contribute to an increased level of standardization of proteomic procedures by two ways. The filmed protocol is used to increase the level of details that can be described precisely. Moreover, the availability of standardized procedures to process biological replicates should guarantee a higher robustness of data, taking into account also the human factor within the technical reproducibility of the extraction procedure.
The protocol described requires 16 days for its completion: fourteen days for cultures and two days for protein extraction (figure 1). 
Briefly, Fusarium strains are grown on solid media for 4 days; they are then manually fragmented and transferred into a modified toxin inducing media (Jiao et al., 2008) for 10 days. Mycelium is collected by filtration through a Miracloth layer. Grinding is performed in a cold chamber. Different operators performed extraction replicates (n=3) in order to take into account the bias due to technical variations (figure 2). Extraction was based on a SDS/DTT buffer as described in Taylor et al. (2008) with slight modifications. Total protein extraction required a precipitation process of the proteins using Aceton/TCA/DTT buffer overnight and Acetone /DTT washing (figure 3a,3b). Proteins were finally resolubilized in the protein-labelling buffer and quantified. Results of the extraction were visualized on a 1D gel (Figure 4, SDS-PAGE), before proceeding to 2D gels (IEF/SDS-PAGE). The same procedure can be applied for proteomic analyses on other growing media and other filamentous fungi (Miles et al., 2007).

Toxin Induction and Protein Extraction from Fusarium spp. Cultures for Proteomic Studies

Protein extraction for proteomic analyses in fungal species requires high levels of standardization to be accomplished according with the minimum information about a proteomic experiment (MIAPE) guidelines. We present a video-protocol that includes a procedure for minimizing experimental bias during toxin induction and protein extraction from Fusarium spp.

Fusaria are filamentous fungi able to produce different toxins. Fusarium mycotoxins such as deoxynivalenol, nivalenol, T2, zearelenone, fusaric acid, ...

Evisceration of Mouse Vitreous and Retina for Proteomic Analyses

While the mouse retina has emerged as an important genetic model for inherited retinal disease, the mouse vitreous remains to be explored. The vitreous is a highly aqueous extracellular matrix overlying the retina where intraocular as well as extraocular proteins accumulate during disease.1-3 Abnormal interactions between vitreous and retina underlie several diseases such as retinal detachment, proliferative diabetic retinopathy, uveitis, and proliferative vitreoretinopathy.1,4 The relative mouse vitreous volume is significantly smaller than the human vitreous (Figure 1), since the mouse lens occupies nearly 75% of its eye.5 This has made biochemical studies of mouse vitreous challenging. In this video article, we present a technique to dissect and isolate the mouse vitreous from the retina, which will allow use of transgenic mouse models to more clearly define the role of this extracellular matrix in the development of vitreoretinal diseases.

The dissection technique illustrates evisceration of the vitreous, retina, and lens from the mouse eye, separation by centrifugation, and characterization with protein assays.

While the mouse retina has emerged as an important genetic model for inherited retinal disease, the mouse vitreous remains to be explored. The vitreous is ...

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling processes. Extracting sterol-enriched membrane microdomains from plant cells for proteomic analysis is a difficult task mainly due to multiple preparation steps and sources for contaminations from other cellular compartments. The plasma membrane constitutes only about 5-20% of all the membranes in a plant cell, and therefore isolation of highly purified plasma membrane fraction is challenging. A frequently used method involves aqueous two-phase partitioning in polyethylene glycol and dextran, which yields plasma membrane vesicles with a purity of 95% 1. Sterol-rich membrane microdomains within the plasma membrane are insoluble upon treatment with cold nonionic detergents at alkaline pH. This detergent-resistant membrane fraction can be separated from the bulk plasma membrane by ultracentrifugation in a sucrose gradient 2. Subsequently, proteins can be extracted from the low density band of the sucrose gradient by methanol/chloroform precipitation. Extracted protein will then be trypsin digested, desalted and finally analyzed by LC-MS/MS. Our extraction protocol for sterol-rich microdomains is optimized for the preparation of clean detergent-resistant membrane fractions from Arabidopsis thaliana cell cultures.
We use full metabolic labeling of Arabidopsis thaliana suspension cell cultures with K15NO3 as the only nitrogen source for quantitative comparative proteomic studies following biological treatment of interest 3. By mixing equal ratios of labeled and unlabeled cell cultures for joint protein extraction the influence of preparation steps on final quantitative result is kept at a minimum. Also loss of material during extraction will affect both control and treatment samples in the same way, and therefore the ratio of light and heave peptide will remain constant. In the proposed method either labeled or unlabeled cell culture undergoes a biological treatment, while the other serves as control 4.

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Here we describe a robust method for the fractionation of plant plasma membranes into detergent resistant and detergent soluble membranes based on a mixture of unlabeled and in vivo fully 15N labeled Arabidopsis thaliana cell cultures. The procedure is applied for comparative proteomic studies to understand signaling processes.

Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling ...