This work was funded by the Natural Sciences and Engineering Research Council of Canada. The authors thank Bioinformatics Solutions Inc. (Waterloo, Canada) and SPARC BioCentre (Molecular Analysis) at the Hospital for Sick Children (Toronto, Canada) for their contributions to the acquisition of MS data.

1. Material considerations and sample pre-preparation
<ol>
	<li>Use only high purity solvents (acetone, chloroform, methanol) (&#62;99.5%) and chemicals, free of excess moisture.</li>
	<li>Prepare sodium chloride and zinc sulfate solutions (1 M) in water. 
	NOTE: Salt solutions can be stored indefinitely at room temperature, as&#160;long as they are free of contaminant or microbial growth.</li>
	<li>Use the smallest polypropylene (PP) microcentrifuge vial sufficient to retain the required volume of sample and solvents to induce precipitation.</li>
	<li>Ensure that the SDS concentration in the sample to be precipitated is no greater than 2% (w/v). If SDS is higher, dilute the sample with water.</li>
	<li>Ensure a protein concentration between 0.01 and 10 g/L for optimal precipitation efficiency. 
	NOTE: The optimal mass for precipitation ranges between 1-100 &#181;g of protein.</li>
	<li>Ensure all solvents and solutions are free of particulate matter before use. Perform either a filtration (&#60;0.5 &#181;m) or centrifugation step (10,000 x g for 1 min at room temperature) to remove undissolved particulates.</li>
	<li>If disulfide bond reduction and alkylation are required, conduct these steps prior to protein precipitation. Excess reducing and alkylating reagents will be removed through the precipitation process.</li>
	<li>Precipitate the proteins by selecting and performing one of the protocols (steps 2, 3, 4, or 5).</li>
</ol>
2. Rapid (vial-based) protein precipitation with acetone
<ol>
	<li>Pipet 90 &#181;L of (particulate-free) protein or proteome solution into a PP microcentrifuge tube. Then, add 10 &#181;L of 1 M aqueous NaCl. 
	NOTE: If the ionic strength of the proteome extract already exceeds 100 mM, no additional salt is necessary.</li>
	<li>Pipet 400 &#181;L of acetone into the sample. Cap the vial and tap the vial gently to combine the solvents. Vigorous mixing is not required. 
	NOTE: The volume of protein, salt, and acetone can be increased so long as the relative ratio of each is maintained.</li>
	<li>Allow the vial to incubate at room temperature, undisturbed, for a minimum of 2 min. 
	NOTE: Longer incubations, including those at reduced temperature (e.g., conventional acetone precipitation employs overnight precipitation in the freezer), may result in the formation of larger (visible) aggregated protein particulates (Figure 1A), which generally do not improve total protein recovery.</li>
	<li>Following incubation, place samples in a centrifuge, noting the orientation of the vial. Spin for a minimum of 2 min, at 10,000 x g or higher at room temperature.</li>
	<li>Uncap the vial and gently decant the supernatant by slowly inverting the vial to a waste container. Touch the inverted vial to a paper towel to draw residual solvent from the vial. 
	CAUTION: Waste solvents should be retained and discarded as per appropriate protocols.</li>
	<li>For SDS-containing samples, dispense 400 &#181;L of fresh acetone, being careful not to disturb the pellet. 
	NOTE: Step 2.6 is optional.
	<ol>
		<li>Immediately centrifuge the sample (10,000 x g or higher for 1 min at room temperature), placing the vial into the rotor in the same orientation as the initial spin. Decant the wash solvent as described in step 2.5.</li>
	</ol>
	</li>
	<li>Allow the sample to fully dry with the cap open (~1 min). Recap the vial and proceed with pellet solubilization (step 6).</li>
</ol>
3. Precipitation of low molecular weight (LMW) peptides (ZnSO4 + acetone)
<ol>
	<li>Dispense 54 &#181;L of proteome extract to a 2 mL PP vial, and then add 6 &#181;L of 1 M ZnSO4. 
	NOTE: Optimal recovery of LMW peptides (&#8804;5 kDa) is obtained by adding acetone to a final 97% by volume. Assuming a 2 mL PP vial, the maximal initial sample volume is 54 &#181;L.</li>
	<li>Add 1940 &#181;L of acetone to a final 97% by volume. Swirl gently to mix, and let stand undisturbed on the benchtop for a minimum of 2 min.</li>
	<li>Centrifuge (10,000 x g for 1 min at room temperature) and remove the supernatant by inverting the vial, and then touching the vial to a paper towel.</li>
	<li>For SDS-containing samples, dispense 400 &#181;L of fresh acetone, being careful not to disturb the pellet. 
	NOTE: Step 3.4 is optional.
	<ol>
		<li>Immediately centrifuge the sample as per step 3.3, placing the vial into the rotor in the same orientation as the initial spin. Decant the wash solvent as described in step 3.3.</li>
	</ol>
	</li>
	<li>Re-solubilize the resulting dry pellet in an aqueous solvent with brief vortexing or sonication (~5 min).</li>
</ol>
4. Protein precipitation by chloroform/methanol/water (CMW)
<ol>
	<li>Dispense 100 &#181;L of the protein or proteome solution into a PP vial. Add 400 &#181;L of methanol, followed by 100 &#181;L of chloroform. Cap the vial and vortex briefly to mix. 
	NOTE: For CMW precipitation, 1.5 mL vials with narrow bottoms are preferred (Figure 2A). 
	CAUTION: Chloroform solvent should be handled in an appropriate ventilation hood. All solvents that contact chloroform should be treated as halogenated waste when disposed of.</li>
	<li>Quickly dispense 300 &#181;L of water directly into the center of the vial. Cap the vial. Allow the sample to sit on the benchtop undisturbed for 1 min. 
	NOTE: The solution will immediately appear cloudy white. Avoid mixing the vial following the addition of water.</li>
	<li>Place the PP vial in a centrifuge and spin for a minimum of 5 min (10,000 x g or higher at room temperature). 
	NOTE: Once centrifuged, two visible solvent layers will form (top layer = methanol/water; bottom = chloroform). A solid protein pellet forms at the solvent interface (Figure 2A).</li>
	<li>Using a large (1 mL) micropipette tip, and holding the vial at ~45&#176;, remove ~700 &#181;L of the solvent from the upper layer at a uniform rate.</li>
	<li>Use a smaller (200 &#181;L) micropipette tip to continue removing the upper solvent layer from the ~45&#176; tilted vial. Pipet in one continuous motion until the upper solvent layer forms a bead in the vial.</li>
	<li>Add 400 &#181;L of fresh methanol to the sample vial, without disturbing the pellet, by dispensing the solvent down the side of the vial.</li>
	<li>Cap the vial. Combine the solvent layers by gently rocking the vial to swirl the solvents together. 
	NOTE: It is essential to avoid disrupting the pellet. Do not vortex the vial.</li>
	<li>Noting the orientation of the vial in the rotor, centrifuge for a minimum of 10 min (10,000 x g at room temperature). The protein pellet adheres to the bottom of the vial (Figure 2B).</li>
	<li>Tip the vial at 45&#176;, with the pellet facing down. Place the pipette tip along the upper edge of the vial and remove the supernatant with a 1 mL micropipette tip at a slow but continuous rate. Retain ~20 &#181;L of solvent in the vial.</li>
	<li>Wash the protein pellet for SDS-containing samples by slowly dispensing 400 &#181;L of fresh methanol. Do not vortex the vial.
	<ol>
		<li>Proceed directly with centrifugation (10,000 x g for 2 min at temperature), placing the vial into the rotor with the same orientation as the initial spin.</li>
	</ol>
	</li>
	<li>Remove the solvent, as per step 4.9. Allow the sample to air dry in a fumehood until the residual solvent evaporates.</li>
	<li>Consult the recommended resolubilization procedures in step 6.</li>
</ol>
5. Protein precipitation using a disposable filtration cartridge
NOTE: Each solvent-based precipitation protocol described in steps 2-5 can be performed in a two-stage filtration and extraction cartridge (see Table of Materials).
<ol>
	<li>With the plug attached to the upper filtration cartridge (Figure 3A), dispense the desired volume of the extracted proteome, salt, and solvent as outlined in one of the three options below.
	<ol>
		<li>(Option 1) For protein precipitation with acetone, combine 90 &#181;L of protein or proteome solution, 10 &#181;L of 1 M aqueous NaCl, and 400 &#181;L of acetone. Incubate for a minimum of 2 min on the benchtop. 
		NOTE: A visible precipitate will develop for concentrated protein samples (1 g/L) (Figure 3B).</li>
		<li>(Option 2) For LMW peptide precipitation, combine 15 &#181;L of the sample, 1.5 &#181;L of 1 M ZnSO4, and 485 &#181;L of acetone. Incubate for a minimum of 2 min on the benchtop. 
		NOTE: A salt concentration of 90 mM in the aqueous sample will not impact recovery relative to the 100 mM recommended in step 3.</li>
		<li>(Option 3) For CMW precipitation, add 50 &#181;L of proteome extract, 200 &#181;L of methanol, and 50 &#181;L of chloroform. Cap the vial and briefly vortex to combine.
		<ol>
			<li>Quickly dispense 150 &#181;L of water directly into the center of the vial. Incubate for 1 min on the benchtop.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Centrifuge for 2 min at 2,500 x g at room temperature with the plug still attached to the filtration cartridge.</li>
	<li>Invert the cartridge, and then unscrew and remove the plug from the cartridge base.</li>
	<li>Place the filtration cartridge in a clean vial and return to the centrifuge. Spin for 3 min at 500 x g at room temperature. Discard the flow-through solvent from the lower vial. 
	NOTE: If any solvent remains in the upper filtration cartridge, return to the centrifuge and perform an additional spin.</li>
	<li>Wash the protein pellet by adding 400 &#181;L of acetone to the filtration cartridge (for CMW precipitation, step 5.1.3, add 400 &#181;L of methanol).</li>
	<li>Centrifuge for 3 min at 500 x g at room temperature or until no solvent remains in the upper cartridge.</li>
	<li>Re-solubilize the precipitation pellet as described in step 6.</li>
</ol>
6. Resolubilization of protein pellet
<ol>
	<li>Wet the membrane at the base of the filtration cartridge by dispensing 2-5 &#181;L of isopropanol directly to the membrane immediately before the resolubilization protocols described below.</li>
	<li>Follow one of the following resolubilization methods.
	<ol>
		<li>(Option 1) Add a minimum of 20 &#181;L of aqueous buffer containing &#8805;2% SDS to the filtration cartridge. Cap and vortex vigorously (~1 min). Alternatively, sonicate (&#62;10 min) to disperse the protein pellet.
		<ol>
			<li>Heat the sample at 95 &#176;C for 5 min). Repeat the mixing step after heating. 
			NOTE: Laemmli gel loading buffer can re-solubilize the protein pellet. However, SDS-containing samples are incompatible with trypsin digestion and reversed-phase LC and MS.</li>
		</ol>
		</li>
		<li>(Option 2) Prepare a solution of 80% (v/v) formic acid in water. Prechill the acid solution (-20 &#176;C), as well as the filtration cartridge containing precipitated protein.
		<ol>
			<li>Dispense 50 &#181;L of cold formic acid into the cartridge; cap and vortex for 30 s. Return to the freezer (-20 &#176;C) for 10 min.</li>
			<li>Vortex the cartridge again for 30 s. Then, repeat the chilling and mixing cycle one more time (10 min, -20 &#176;C, 30 s vortex).</li>
			<li>Add water to a final 500 &#181;L, diluting the formic acid to 8%. 
			NOTE: The cold formic acid protocol is incompatible with subsequent trypsin digestion but is compatible with LC-MS.</li>
		</ol>
		</li>
		<li>(Option 3) Add 50 &#181;L of freshly prepared 8 M urea in water to the filtration cartridge. Sonicate for 30 min.
		<ol>
			<li>Allow the cartridge to incubate on the benchtop for 1 h (up to overnight).</li>
			<li>Dilute the 8 M urea a minimum 5-fold with water or appropriate buffer. 
			NOTE: Once diluted, the urea solubilization protocol is compatible with subsequent trypsin digestion, as well as LC-MS.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
7. Protein digestion
<ol>
	<li>For bottom-up MS analysis, subject the re-solubilized proteins to enzymatic digestion using one of the two methods mentioned below.
	<ol>
		<li>(Option 1) For formic acid resolubilization, reduce the initial volume of 80% formic acid in step 6.2.2.1 to 25 &#181;L. In step 6.2.2.3, use 375 &#181;L of water to dilute the formic acid to 5% (v/v).</li>
		<li>Dispense pepsin into the cartridge at an approximate protein to enzyme ratio of 50:1. With a plug attached to the filtration cartridge, incubate the sample overnight at room temperature.</li>
	</ol>
	</li>
	<li>(Option 2) For resolubilization in urea, ensure a pH between 8 and 8.3 with the inclusion of 100 mM of Tris or ammonium bicarbonate in step 6.2.3.2.
	<ol>
		<li>Add trypsin at an approximate protein to enzyme mass ratio of 50:1. With a plug attached to the cartridge, incubate the sample overnight in a warm water bath at 37 &#176;C.</li>
		<li>Terminate the digestion by acidifying the solution with 10% TFA to a final 1%.</li>
	</ol>
	</li>
	<li>Recover the pepsin- or trypsin-digested protein by removing the plug from the base of the filter and centrifuging the cartridge contained within a clean vial (2 min, 5000 x g, room temperature).</li>
</ol>
8. SPE clean-up
NOTE: For additional sample desalting following digestion or solvent exchange, the sample can be subject to reversed-phase clean-up as described.
<ol>
	<li>Prime an SPE cartridge (see Table of Materials) by passing 300 &#181;L of methanol (2 min, 400 x g) followed by 300 &#181;L of 5% acetonitrile/0.1% of TFA (2 min, 400 x g).</li>
	<li>Connect the primed SPE cartridge to the base of the filtration cartridge containing re-solubilized or digested protein.</li>
	<li>Spin the protein through the SPE cartridge (5 min, 800 x g at room temperature). If solvent remains in the upper cartridge, return the cartridge to the centrifuge and repeat the spin. 
	NOTE: Passing the sample through the SPE cartridge a second time may improve recovery.</li>
	<li>Add 300 &#181;L of 5% acetonitrile/0.1% of TFA in water to the cartridge. To wash, flow through the SPE cartridge (2 min, 2000 x g). Discard the flow-through.</li>
	<li>For LMW proteins or digested peptides, elute the sample by flowing 300 &#181;L of 50% acetonitrile/0.1%&#160;TFA (5 min, 2500 x g).</li>
	<li>For intact proteins, follow step 8.5 with an additional elution step using 300 &#181;L of 75% acetonitrile/0.1% TFA. Combine the two resulting extracts. 
	NOTE: Step 8.6 is optional.</li>
</ol>

The Doucette laboratory conceived of and patented the ProTrap XG employed in this study. AAD is also a founding partner of Proteoform Scientific, which commercialized the sample preparation cartridge.

Optimal MS characterization is achieved when residual SDS is depleted below 10 ppm. While alternative approaches, such as FASP and on-bead digestion, offer quantitative SDS depletion with variable recovery31,32,33, the primary objective of precipitation is to maximize purity and yield simultaneously. This depends on effectively isolating the supernatant (containing the SDS) without disturbing the protein pellet. With vial-based precipitation, once the bulk of the supernatant is removed by pipetting, it is increasingly likely that some of the aggregated pellets are accidentally lost. For this reason, it is essential to leave behind a more significant fraction of the residual solvent (~20 &#181;L) and to add a washing step34. The washing step dilutes and removes the residual solvent from the vial. Particularly with CMW, it is unnecessary to vortex the sample vial once the pellet has formed. Disrupting the pellet through vigorous agitation has the unwanted effect of increasing the likelihood of loss from accidental pipetting. If vortexing is included (as recommended by previous protocols)35, the potential exists for portions of the CMW pellet to adhere to the underside of the vial cap; once centrifuged, the pellet remains fixed on the vial cap and can result in ~50% loss.
Rapid precipitation can be performed with high recovery of dilute proteome samples, ideally between 0.01-2 mg/mL, or a corresponding protein mass between 1-200 &#181;g. However, quantitative and reproducible recoveries starting from below 0.01 mg/mL protein may benefit from longer precipitation times ranging from 10 min to 1 h, demonstrating throughput limitations of the precipitation workflow. Surprisingly, more concentrated samples (10 mg/mL) show a statistical reduction in yield, presumably from accidental pipetting losses. Assuming &#62;10 &#181;g protein, a visible pellet should be observed on the side of the vial (Figure 1B). Smaller quantities, down to 1 &#181;g, are challenging to see. This challenges the capacity to pipette the supernatant without disrupting the protein pellet. The vial can be inverted (slowly) with acetone to separate the solvent from the pellet. For CMW, the pellet does not reliably adhere sufficiently to the vial, thereby favoring pipetting over decanting of the supernatant. For vial-based precipitation, working with the smallest possible microcentrifuge tube is recommended to facilitate the intended sample and solvent volumes. Precipitation in the disposable filtration cartridge employed in this work provides a maximum volume capacity of 500 &#181;L, enabling protein precipitation with 80% acetone on sample volumes up to 100 &#181;L. Sample, salt, and solvent volumes can be adjusted accordingly if the recommended concentrations are maintained.
The purity of the protein pellet recovered from organic solvent-based precipitation is limited by the complexity of the sample matrix, buffer components, and precipitation conditions. For example, specific buffer components such as glycine (used for SDS PAGE separations) have been shown to co-precipitate with protein using 80% acetone. However, glycine remains soluble through CMW precipitation. Acetone has been reported to precipitate DNA fragments36,37, potentially adding undesired background impurities to the recovered pellet. Precipitation of low molecular weight proteins and peptides requires an elevated level of organic solvent and a specific salt type to maximize yield. While several salts have been explored, ZnSO4 provides consistently high products. This salt will precipitate in 97% acetone in the absence of protein. Thus, the resulting protein pellet contains a high concentration of salt. It is noted that employing 90% acetone by volume will also achieve high peptide yields, though a statistically significant drop (~5%) in recovery is expected. However, this allows processing a more significant sample volume (up to 180 &#181;L, with 20 &#181;L of 1 M ZnSO4) in each 2 mL vial. Beyond matrix impurities co-precipitating with the protein pellet, it must be stated that solvent precipitation inherently causes denaturation of the sample38,39. Therefore, this protocol is not applicable for the preparations of functional proteins or native MS workflows. Acetone has also been reported to cause covalent protein modifications at glycine residues40&#160;and induce a +98 u mass shift, speculated to be a byproduct of aldol condensation acetone41.
When employing a filtration cartridge for protein precipitation, isolation of the protein pellet relies on the retention of aggregates above a PTFE membrane filter. The porosity of this membrane exceeds that of a molecular weight cutoff filter (as seen in FASP), permitting protein isolation with reduced spin times. Rapid solution transfer at low spin speeds relies on proper wetting of the PTFE membrane; organic solvents readily flow through, though a dry PTFE filter impedes aqueous solvents. If the filtration cartridge appears to be clogged, the membrane should be re-wetted by directly applying a small volume of organic solvent (e.g., isopropanol) to the filter. Depending on the size of the protein pellet and the volume of sample employed, additional centrifugation or spins at higher speeds (up to 3,000 x g) may be required to ensure all solvent has passed through the filtration cartridge.
Protein recovery from precipitation with optimal conditions is ultimately limited by the challenge of pellet re-solubilization, with few solvent options being compatible with downstream processing and LC-MS. Additionally, several precipitation conditions such as long exposures to low temperatures and over-drying a tightly packed pellet contribute to re-solubilization challenges40. It is noted that CMW protein pellets are generally less soluble than acetone pellets. Maximized re-solubilization efficiency of precipitated protein by 80% cold formic acid (step 6.2.2) has previously been reported42; the cold temperature prevents protein modification, which otherwise occurs in concentrated formic acid43,44. Diluting the acid concentration also slows the modification reaction. Formic acid is recommended for top-down MS approaches or before enzymatic digestion with pepsin. Employing this solvent demands little physical treatment; the addition of only 5 &#181;L (enough to cover the protein pellet) may be sufficient when combined with vortexing, brief sonication, or repeat pipetting. Similarly, for samples intended to be analyzed by SDS PAGE, re-dissolving in SDS-containing Laemmli buffer is highly effective, when combined with modest mixing of the sample prior to heating. However, these solvents are both incompatible with trypsin. Resolubilization with 8 M urea is recommended prior to trypsin digestion, ensuring that the urea has been freshly prepared (same day). A minimum volume of 50 &#181;L buffer is recommended for protein re-solubilization within the filtration cartridge to maximize contact between the chaotropic solvent and pellet, as well as to aid dissolution during sonication, repeat pipetting and/or vortexing. Alternative approaches exploit trypsin to re-solubilize the protein, meaning the protein need not be fully re-dissolved prior to enzyme addition. However, this approach can bias digestion, favoring the more soluble species while hydrophobic proteins experience shorter digestion time45. Addition of 8 M urea, together with basic buffers such as Tris or ammonium bicarbonate, demands a post-digestion sample clean-up step. For such sample additives, reversed phase column clean-up is ideal. The disposable filtration cartridge employed in this study is supplemented with an interchangeable reversed phase SPE cartridge. This cartridge is also ideally suited for solvent exchange, in the case of the formic acid resolubilization protocol. It is important to note that any solid phase extraction approach is associated with inherent loss in sample recovery. Therefore, the user should weigh the benefits of recovery and the additional purification for their experiment.
It is anticipated that these protocols will enable proteomics researchers to streamline their detergent-based workflows, capitalizing on SDS for proteome extraction. Preparative strategies that facilitate consistent recovery of the complete proteome are critical. A two-stage spin cartridge simplifies the opportunity for rapid, robust, and reproducible proteome sample isolation. Such an approach would be amenable to applications requiring rigorous analysis without sacrificing sample throughputs, such as clinical settings or large-scale research initiatives46. Future applications of these approaches may include biomarker discovery, detection, accurate quantitation, and drug and drug target discovery.

Proteome analysis by mass spectrometry has become increasingly rigorous, owing to the enhanced sensitivity, resolution, scan speed, and versatility of modern MS instruments. MS advances contribute to greater protein identification efficiency and more precise quantitation1,2,3,4,5. With improved MS instrumentation, researchers demand a correspondingly consistent front-end sample preparation strategy capable of quantitative recovery of high-purity proteins in minimal time across all stages of the workflow6,7,8,9,10,11. To accurately reflect the proteome status of a biological system, proteins must be isolated from the native sample matrix in an efficient and unbiased fashion. To this end, including a denaturing surfactant, such as sodium dodecyl sulfate (SDS), ensures efficient protein extraction and solubilization12. However, SDS strongly interferes with electrospray ionization, causing severe MS signal suppression if not properly eliminated13.
Various SDS depletion strategies are available for subsequent proteome analysis, such as the retention of proteins above a molecular weight cutoff filter contained within disposable spin cartridges14,15,16. The filter-aided sample preparation method (FASP) is favored as it effectively depletes SDS below 10 ppm, facilitating optimal MS. However, protein recovery with FASP is variable, which prompted the exploration of other techniques. Chromatographic approaches that selectively capture protein (or surfactant) have evolved into various convenient cartridges or bead-based formats17,18,19,20,21. Given these simple and (ideally) consistent strategies to protein purification, the classical approach of protein precipitation with organic solvents is often overlooked as a promising approach to protein isolation. While solvent precipitation is shown to deplete SDS below critical levels successfully, protein recovery has been a longstanding concern of this approach. Multiple groups have observed a protein recovery bias, with unacceptably low precipitation yields as a function of protein concentration, molecular weight, and hydrophobicity22,23. Due to the diversity of precipitation protocols reported in the literature, standardized precipitation conditions were developed. In 2013, Crowell et al. first reported the dependence of ionic strength on the precipitation efficiency of proteins in 80% acetone24. For all proteins examined, the addition of up to 30 mM sodium chloride was shown to be essential to maximize yields (up to 100% recovery). More recently, Nickerson et al. showed that the combination of even higher ionic strength (up to 100 mM) with elevated temperature (20 &#176;C) during acetone precipitation gave near quantitative recovery in 2-5 min25. A slight drop in the recovery of low molecular weight (LMW) proteins was observed. Therefore, a subsequent report by Baghalabadi et al. demonstrated the successful recovery of LMW proteins and peptides (&#8804;5 kDa) by combining specific salts, particularly zinc sulfate, with a higher level of organic solvent (97% acetone)26.
While refining the precipitation protocol lends a more reliable protein purification strategy for MS-based proteomics, the success of conventional precipitation relies heavily on user technique. A primary goal of this work is to present a robust precipitation strategy that facilitates the isolation of the protein pellet from the contaminating supernatant. A disposable filtration cartridge was developed to eliminate pipetting by isolating aggregated protein above a porous PTFE membrane filter27. MS-interfering components in the supernatant are effectively removed in a short, low-speed&#160;centrifugation step. The disposable filter cartridge also offers an interchangeable SPE cartridge, which facilitates subsequent sample clean-up following resolubilization and optional protein digestion, ahead of mass spectrometry.
A series of recommended proteome precipitation workflows are presented here, including modified acetone and chloroform/methanol/water28 protocols, in a conventional (vial-based) and a semi-automated format in a disposable two-state filtration and extraction cartridge. The resulting protein recoveries and SDS depletion efficiencies are highlighted, together with bottom-up LC-MS/MS proteome coverage, to demonstrate the expected outcome from each protocol. The practical benefits and drawbacks associated with each approach are discussed.

<table><tbody><tr><td>Acetone</td><td>Fisher Scientific</td><td>AC177170010</td><td>&amp;le;0.002 % aldehyde</td></tr><tr><td>Acetonitrile</td><td>Fisher Scientific</td><td>A998-4</td><td>HPLC grade</td></tr><tr><td>Ammonium Bicarbonate</td><td>Millipore Sigma</td><td>A6141-1KG</td><td>solid</td></tr><tr><td>Beta mercaptoethanol</td><td>Millipore Sigma</td><td>M3148-25ML</td><td>Molecular biology grade</td></tr><tr><td>Bromophenol blue</td><td>Millipore Sigma</td><td>B8026-5G</td><td>Bromophenol blue sodium salt</td></tr><tr><td>Chloroform</td><td>Fisher Scientific</td><td>C298-400</td><td>Chloroform</td></tr><tr><td>Formic Acid</td><td>Honeywell</td><td>56302</td><td>Eluent additive for LC-MS</td></tr><tr><td>Fusion Lumos Mass Spectrometer</td><td>ThermoFisher Scientific</td><td></td><td>for analysis of standard protein mixture</td></tr><tr><td>Glycerol</td><td>Millipore Sigma</td><td>356352-1L-M</td><td>For molecular biology, &gt; 99%</td></tr><tr><td>Isopropanol</td><td>Fisher Scientific</td><td>A4641</td><td>HPLC grade</td></tr><tr><td>Methanol</td><td>Fisher Scientific</td><td>A452SK-4</td><td>HPLC grade</td></tr><tr><td>Microcentrifuge</td><td>Fisher Scientific</td><td>75-400-102</td><td>up to 21,000 xg</td></tr><tr><td>Microcentrifuge Tube (1.5 mL)</td><td>Fisher Scientific</td><td>05-408-130</td><td>tapered bottom</td></tr><tr><td>Microcentrifuge Tube&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; (2 mL)</td><td>Fisher Scientific</td><td>02-681-321</td><td>rounded bottom</td></tr><tr><td>Micropipette Tips&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; (0.1-10 &amp;mu;L)</td><td>Fisher Scientific</td><td>21-197-28</td><td>Universal pipet tip, non-sterile</td></tr><tr><td>Micropipette Tips&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; (1-200 &amp;mu;L)</td><td>Fisher Scientific</td><td>07-200-302</td><td>Universal pipet tip, non-sterile</td></tr><tr><td>Micropipette Tips&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; (200-1000 &amp;mu;L)</td><td>Fisher Scientific</td><td>07-200-303</td><td>Universal pipet tip, non-sterile</td></tr><tr><td>Micropipettes</td><td>Fisher Scientific</td><td>13-710-903</td><td>Micropipet Trio pack</td></tr><tr><td>Pepsin</td><td>Millipore Sigma</td><td>P0525000</td><td>Lyophilized powder,&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &gt;3200 units/ mg</td></tr><tr><td>ProTrap XG</td><td>Proteoform Scientific</td><td>PXG-0002</td><td>50 complete units per box</td></tr><tr><td>Sodium Chloride</td><td>Millipore Sigma</td><td>S9888-1KG</td><td>ACS reagent, &gt;99 %</td></tr><tr><td>Sodium Dodecyl Sulfate</td><td>ThermoFisher Scientific</td><td>28312</td><td>powdered solid</td></tr><tr><td>timsTOF Pro Mass Spectrometer</td><td>Bruker</td><td></td><td>for analysis of liver proteome extract</td></tr><tr><td>Trifluoroacetic Acid</td><td>ThermoFisher Scientific</td><td>L06374.AP</td><td>99%</td></tr><tr><td>Tris</td><td>Fisher Scientific</td><td>BP152-500</td><td>Molecular biology grade</td></tr><tr><td>Trypsin</td><td>Millipore Sigma</td><td>9002-07-7</td><td>From bovine pancreas, TPCK-treated</td></tr><tr><td>Urea</td><td>Bio-Rad</td><td>1610731</td><td>solid</td></tr><tr><td>Water (deionized)</td><td>Sartorius Arium Mini Water Purification System</td><td>76307-662</td><td>Type 1 ultrapure (18.2 M&amp;Omega; cm)</td></tr><tr><td>Zinc Sulfate</td><td>Millipore Sigma</td><td>307491-100G</td><td>solid</td></tr></tbody></table>

organic solvent-based protein precipitation for robust proteome purification ahead of mass spectrometry

While multiple advances in mass spectrometry (MS) instruments have improved qualitative and quantitative proteome analysis, more reliable front-end approaches to isolate, enrich, and process proteins ahead of MS are critical for successful proteome characterization. Low, inconsistent protein recovery and residual impurities such as surfactants are detrimental to MS analysis. Protein precipitation is often considered unreliable, time-consuming, and technically challenging to perform compared to other sample preparation strategies. These concerns are overcome by employing optimal protein precipitation protocols. For acetone precipitation, the combination of specific salts, temperature control, solvent composition, and precipitation time is critical, while the efficiency of chloroform/methanol/water precipitation depends on proper pipetting and vial manipulation. Alternatively, these precipitation protocols are streamlined and semi-automated within a disposable spin cartridge. The expected outcomes of solvent-based protein precipitation in the conventional format and using a disposable, two-stage filtration and extraction cartridge are illustrated in this work. This includes the detailed characterization of proteomic mixtures by bottom-up LC-MS/MS analysis. The superior performance of SDS-based workflows is also demonstrated relative to non-contaminated protein.

Figure 4 summarizes the expected SDS depletion following vial-based or cartridge-facilitated precipitation of proteins in a disposable filter cartridge using acetone. Conventional overnight incubation (-20 °C) in acetone is compared to the rapid acetone precipitation protocol at room temperature (step 2), as well as CMW precipitation (step 4). Residual SDS was quantified by the methylene blue active substances (MBAS) assay29. Briefly, 100 µL sample was combined with 100 µL MBAS reagent (250 mg methylene blue, 50 g sodium sulfate, 10 mL sulfuric acid, diluted in water to 1.0 L), followed by the addition of 400 µL chloroform and absorbance measurement of the organic layer at 651 nm on a UV/Vis spectrophotometer. All approaches reduce SDS to permit optimal MS analysis.
Quantitative and reproducible protein recovery is achieved following rapid acetone precipitation and CMW precipitation, as seen in Figure 5 through SDS PAGE analysis of a processed yeast total cell lysate. Precipitation in a disposable filtration cartridge eliminates the need to carefully pipet the SDS-containing supernatant while retaining the aggregated proteins above a membrane filter. Consistent recovery is obtained with all precipitation protocols, with no visible bands detected in the supernatant fractions across three independent replicates.
Figure 6 quantifies the expected yields, including the resolubilization of precipitated protein pellets using cold formic acid (step 6). CMW precipitation affords quantitative recovery by carefully preserving the pellet in a vial-based approach (step 5), which equals that obtained using the cartridge (100 ± 4% vs. 101 ± 3%, respectively). Recovery of acetone-precipitated protein pellets benefits from a filtration cartridge, with a 15-20 % improvement in yield observed. In vials, isolation of the acetone supernatant from the aggregated protein essentially relies on the adherence of the pellet to the PP tube surface; the filtration cartridge eliminates this concern as the filter ensures high recovery of precipitated protein without pipetting.
To efficiently recover LMW proteins and peptides, the acetone precipitation protocol is modified by substituting NaCl for ZnSO4 and raising the solvent percentage to 97%. Combining this specific salt and higher levels of organic solvent are required for the high recovery of LMW proteins and peptides26. As seen in Figure 7, cartridge-based protein precipitation demonstrates superior recovery of a pepsin-digested sample of bovine plasma relative to vial-based precipitation. The disposable spin cartridge can recover over 90% of LMW peptides. More significant differences in yield are noted in the cartridge when employing NaCl, confirming the importance of salt type to maximize yield. Including ZnSO4 as opposed to NaCl results in an aggregated protein pellet that is more readily trapped by the spin cartridge filter.
To assess the efficacy of precipitating proteins over a wide dynamic range, a mixture of three standard proteins was processed: β-galactosidase (β-gal) from E. coli, cytochrome c (Cyt c) from bovine, and enolase (Eno) from S. cerevisiae. The mass ratio of β-gal:Cyt c:Eno was 10,000:10:1. Samples initially contained 2% of SDS prior to cartridge-based precipitation (step 5) and were re-solubilized and digested with trypsin (steps 6 and 7). Samples prepared in vials acted as a control, having no SDS and omitting the precipitation. All samples were subject to equivalent SPE clean-up (step 8). Bottom-up MS was conducted, with MS/MS spectra searched against a combined database containing all proteins from the three species involved (see Table of Materials for instrument and software platforms). A peptide false discovery rate of 1% was employed. All three proteins were identified by MS, with 666, 28, and 35 unique peptides for β-gal, Cyt c, and Eno, respectively. Figure 8 quantifies the relative ratio (peptide peak intensity) from each sample, with a ratio above 1 reflecting a higher peptide abundance for samples processed in the disposable filter cartridges. The results demonstrate the benefits of incorporating SDS into a proteomics workflow, minimizing protein loss (e.g., from potential adsorption to the sample vial), and maximizing peptide yields.
Bovine liver was procured at a local grocery store. The proteins were isolated by extracting the tissue with a solution of 1% SDS. Subsequently, the recovered proteome was precipitated, re-solubilized (urea), and digested with trypsin, all within a disposable cartridge. Bottom-up LC-MS/MS was conducted, resulting in the identification of an average of ~8,000 proteins (~30,000 peptides). False discovery rates of 0.5% and 1.0% for peptide spectra and protein groups, was employed, searching the bovine database. The technical reproducibility of this cartridge-based workflow is assessed through overlapping protein identifications. The replicate MS injections of a common digested sample achieves on average 78 ± 0.5% overlap with the identified proteins. By comparison, samples independently prepared in discrete cartridges achieved 76 ± 0.5% overlap. These data suggest that the contribution of sample preparation toward the total variability of the analysis is minor, relative to that already contributed by the LC-MS instrumental approach. The bovine proteins identified from three technical replicates (processed independently in three disposable cartridges) were further characterized concerning their molecular weight, hydrophobicity, and isoelectric point, shown in Figure 9. A two-way ANOVA could not determine statistical differences in the identified proteomes across the technical replicates. Finally, Figure 10 compares the number of identified peptides per protein across the three replicate sample preparations. The correlation coefficients in these graphs (0.94-0.95) demonstrate the high consistency of the sample preparation approach for bottom-up MS analysis.
<img alt="Centrifuge tubes with phase separation in protein purification experiment; image compares conditions A and B." class="xfigimg" src="/files/ftp_upload/63503/63503fig01.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Acetone-precipitated proteins. Samples containing 100 and 1,000 µg of protein combined with 100 mM NaCl and precipitated with 80% acetone (A) following 5 min precipitation time and (B) following precipitation and subsequent centrifugation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63503/63503fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Centrifugation process in microcentrifuge tubes; separation and precipitation of biological samples." class="xfigimg" src="/files/ftp_upload/63503/63503fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Protein precipitation by chloroform/methanol/water. A sample containing 50 µg of protein precipitated as per step 4. (A) Immediately following step 4.4. (B) Immediately following step 4.8. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63503/63503fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Protein aggregation in centrifuge tubes; experimental setup shows aggregation labeled; protein study." class="xfigimg" src="/files/ftp_upload/63503/63503fig03.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: Photos of a disposable two-stage filtration and extraction cartridge for protein precipitation. A sample containing 100 µg protein was combined with 100 mM of NaCl and 80% acetone in (A) the assembled filtration and SPE cartridge and (B) precipitated for 5 min until protein aggregates became visible. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63503/63503fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Precipitation approach results; bar chart of % SDS depleted using vial and spin cartridge methods." class="xfigimg" src="/files/ftp_upload/63503/63503fig04.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4: SDS depletion efficiency following protein precipitation. The percentage of SDS removed is shown from acetone precipitation with the conventional protocol (overnight at -20 °C), the rapid protocol (2 min incubation at room temperature), or by chloroform/methanol/water (CMW) precipitation of an S. cerevisiae lysate, both in conventional (vial) and cartridge format. These samples initially contained 0.5% SDS (5,000 ppm), inferring &gt;99.8% SDS removal is required for optimal MS analysis. Residual SDS is quantified by methylene blue active substances (MBAS) assay. Error bars represent the standard deviation from technical replicates (n = 3). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63503/63503fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Protein precipitation methods comparison; electrophoresis results; acetone, chloroform/methanol." class="xfigimg" src="/files/ftp_upload/63503/63503fig05.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
Figure 5: Total proteome recovery through precipitation. SDS PAGE shows the recovery of S. cerevisiae total protein lysate, precipitated by (A) conventional acetone precipitation, (B) chloroform/methanol/water precipitation, and (C) rapid acetone precipitation. Protein bands are exclusively observed in the pellet fraction, with no visible bands in the supernatant (Super.). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63503/63503fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Precipitation recovery graph comparing acetone approaches; vial vs spin cartridge methods." class="xfigimg" src="/files/ftp_upload/63503/63503fig06.jpg" data-altupdatedat="1747911167000" data-alt="Figure 6"> 
Figure 6: Superior protein recovery within a filtration cartridge. For precipitation of the S. cerevisiae total protein lysate, the disposable spin cartridge facilitates quantitative recovery with acetone and CMW precipitation. High recovery is also possible with vial-based precipitation, though careful sample manipulation and pipetting are required. LC-UV assessed protein recovery following resolubilization of the pellet with cold formic acid are shown here. Error bars represent the standard deviation from technical replicates (n = 3). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63503/63503fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Protein precipitation efficiency bar chart, acetone conditions, vial vs. spin cartridge recovery." class="xfigimg" src="/files/ftp_upload/63503/63503fig07.jpg" data-altupdatedat="1747911167000" data-alt="Figure 7"> 
Figure 7: High precipitation yields for low molecular weight peptides. A modified acetone precipitation protocol for peptides and proteins ≤5 kDa involves coupling 100 mM of ZnSO4 with 97% acetone to achieve the highest yields. Precipitation facilitated by a disposable filtration cartridge demonstrates improved recovery compared to conventional vial-based precipitation across all three precipitation conditions. Error bars represent the standard deviation from technical replicates (n = 3). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63503/63503fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Box plot showing peptide peak areas for β-galactosidase, cytochrome c, enolase; spin cartridge control." class="xfigimg" src="/files/ftp_upload/63503/63503fig8v2.jpg" data-altupdatedat="1747911167000" data-alt="Figure 8"> 
Figure 8: Higher recovery of standard proteins in SDS-based workflow. Tukey Box-and-Whisker plots30 of relative MS signal intensity for peptides recovered from SDS-containing proteins processed in a disposable filtration cartridge relative to a control sample (no SDS, no precipitation). The proteins employed span a wide concentration dynamic range-β-galactosidase:cytochrome c:enolase = 10,000:10:1. Each quartile within the boxes contains 25% of the distribution, while error bars encompass 95% of the distribution. Mean is indicated by "+" and median by a horizontal line. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63503/63503fig8v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Box plots of protein properties: (A) Molecular Weight, (B) Hydrophobicity, (C) Isoelectric Point." class="xfigimg" src="/files/ftp_upload/63503/63503fig09.jpg" data-altupdatedat="1747911167000" data-alt="Figure 9"> 
Figure 9: Identified protein distributions from technical replicates. Tukey Box-and-Whisker plots characterize (A) the molecular weight, (B) hydrophobicity, and (C) isoelectric point of proteins identified by bottom-up LC-MS/MS following triplicate preparations of a bovine liver lysate in a two-stage filtration and extraction cartridge. There was no statistical difference in these characteristics by two-way ANOVA (p &lt; 0.05). Each quartile within the boxes contains 25% of the distribution, while error bars encompass 95% of the distribution. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63503/63503fig09large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Correlation of peptide IDs, protein samples comparison graphs A, B, C; linear regression equations." class="xfigimg" src="/files/ftp_upload/63503/63503fig10.jpg" data-altupdatedat="1747911167000" data-alt="Figure 10"> 
Figure 10: Correlation of peptide IDs per protein through the SDS-based preparation workflow across preparative replicates. Analysis of bottom-up proteome reproducibility across (A) samples 1 and 2, (B) samples 2 and 3, and (C) samples 1 and 3 based on the number of peptide MS identifications per protein. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63503/63503fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Organic Solvent-Based Protein Precipitation for Robust Proteome Purification Ahead of Mass Spectrometry at JoVE.com

Organic Solvent-Based Protein Precipitation for Robust Proteome Purification Ahead of Mass Spectrometry

The present protocol describes solvent-based protein precipitation under controlled conditions for robust and rapid recovery and purification of proteome samples prior to mass spectrometry.

While multiple advances in mass spectrometry (MS) instruments have improved qualitative and quantitative proteome analysis, more reliable front-end ...

organic-solvent-based-protein-precipitation-for-robust-proteome

Nanyang Technological University

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Analytical Chemistry

Chemistry

Unit 1

Complexometric Titration, Precipitation Titration, and Gravimetry

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9.9K Views.  Dalhousie University. Deze protocollen zijn ontworpen om het herstel en de zuiverheid van eiwitten te maximaliseren door middel van op oplosmiddelen gebaseerde precipitatie. We gaan je een paar trucs laten zien om neerslag in flacons uit te voeren, evenals een semi-geautomatiseerde aanpak in een spincartridgeformaat. We hebben ook het neerslagprotocol geoptimaliseerd om consistent en snel te zijn.Het hele protocol duurt slechts een paar minuten. Deze protocollen tonen aan dat oplosmiddelprecipitatie van eiw...

Video: Organic Solvent-Based Protein Precipitation for Robust Proteome Purification Ahead of Mass Spectrometry

Quantitative Proteomics Using Reductive Dimethylation for Stable Isotope Labeling

Stable isotope labeling of peptides by reductive dimethylation (ReDi labeling) is a method to accurately quantify protein expression differences between samples using mass spectrometry. ReDi labeling is performed using either regular (light) or deuterated (heavy) forms of formaldehyde and sodium cyanoborohydride to add two methyl groups to each free amine. Here we demonstrate a robust protocol for ReDi labeling and quantitative comparison of complex protein mixtures. Protein samples for comparison are digested into peptides, labeled to carry either light or heavy methyl tags, mixed, and co-analyzed by LC-MS/MS. Relative protein abundances are quantified by comparing the ion chromatogram peak areas of heavy and light labeled versions of the constituent peptide extracted from the full MS spectra. The method described here includes sample preparation by reversed-phase solid phase extraction, on-column ReDi labeling of peptides, peptide fractionation by basic pH reversed-phase (BPRP) chromatography, and StageTip peptide purification. We discuss advantages and limitations of ReDi labeling with respect to other methods for stable isotope incorporation. We highlight novel applications using ReDi labeling as a fast, inexpensive, and accurate method to compare protein abundances in nearly any type of sample.

Stable isotope labeling of peptides by reductive dimethylation (ReDi labeling) is a rapid, inexpensive strategy for accurate mass spectrometry-based quantitative proteomics. Here we demonstrate a robust method for preparation and analysis of protein mixtures using the ReDi approach that can be applied to nearly any sample type.

Stable isotope labeling of peptides by reductive dimethylation (ReDi labeling) is a method to accurately quantify protein expression differences between ...

An Aquatic Microbial Metaproteomics Workflow: From Cells to Tryptic Peptides Suitable for Tandem Mass Spectrometry-based Analysis

Meta-omic technologies such as metagenomics, metatranscriptomics and metaproteomics can aid in the understanding of microbial community structure and metabolism. Although powerful, metagenomics alone can only elucidate functional potential. On the other hand, metaproteomics enables the description of the expressed in situ metabolism and function of a community. Here we describe a protocol for cell lysis, protein and DNA isolation, as well as peptide digestion and extraction from marine microbial cells collected on a cartridge filter unit (such as the Sterivex filter unit) and preserved in an RNA stabilization solution (like RNAlater). In mass spectrometry-based proteomics studies, the identification of peptides and proteins is performed by comparing peptide tandem mass spectra to a database of translated nucleotide sequences. Including the metagenome of a sample in the search database increases the number of peptides and proteins that can be identified from the mass spectra. Hence, in this protocol DNA is isolated from the same filter, which can be used subsequently for metagenomic analysis.

This protocol is for the extraction and concentration of protein and DNA from microbial biomass collected from seawater, followed by the generation of tryptic peptides suitable for tandem mass spectrometry-based proteomic analysis.

Meta-omic technologies such as metagenomics, metatranscriptomics and metaproteomics can aid in the understanding of microbial community structure and ...

Environment

2 in 1: One-step Affinity Purification for the Parallel Analysis of Protein-Protein and Protein-Metabolite Complexes

Cellular processes are regulated by interactions between biological molecules such as proteins, metabolites, and nucleic acids. While the investigation of protein-protein interactions (PPI) is no novelty, experimental approaches aiming to characterize endogenous protein-metabolite interactions (PMI) constitute a rather recent development. Herein, we present a protocol that allows simultaneous characterization of the PPI and PMI of a protein of choice, referred to as bait. Our protocol was optimized for Arabidopsis cell cultures and combines affinity purification (AP) with mass spectrometry (MS)-based protein and metabolite detection. In short, transgenic Arabidopsis lines, expressing bait protein fused to an affinity tag, are first lysed to obtain a native cellular extract. Anti-tag antibodies are used to pull down protein and metabolite partners of the bait protein. The affinity-purified complexes are extracted using a one-step methyl tert-butyl ether (MTBE)/methanol/water method. Whilst metabolites separate into either the polar or the hydrophobic phase, proteins can be found in the pellet. Both metabolites and proteins are then analyzed by ultra-performance liquid chromatography-mass spectrometry (UPLC-MS or UPLC-MS/MS). Empty-vector (EV) control lines are used to exclude false positives. The major advantage of our protocol is that it enables identification of protein and metabolite partners of a target protein in parallel in near-physiological conditions (cellular lysate). The presented method is straightforward, fast, and can be easily adapted to biological systems other than plant cell cultures.

Protein-protein and protein-metabolite interactions are crucial for all cellular functions. Herein, we describe a protocol that allows parallel analysis of these interactions with a protein of choice. Our protocol was optimized for plant cell cultures and combines affinity purification with mass spectrometry-based protein and metabolite detection.

Cellular processes are regulated by interactions between biological molecules such as proteins, metabolites, and nucleic acids. While the investigation of ...

Biochemistry

Dynamic Proteomic and miRNA Analysis of Polysomes from Isolated Mouse Heart After Langendorff Perfusion

Studies in dynamic changes in protein translation require specialized methods. Here we examined changes in newly-synthesized proteins in response to ischemia and reperfusion using the isolated perfused mouse heart coupled with polysome profiling. To further understand the dynamic changes in protein translation, we characterized the mRNAs that were loaded with cytosolic ribosomes (polyribosomes or polysomes) and also recovered mitochondrial polysomes and compared mRNA and protein distribution in the high-efficiency fractions (numerous ribosomes attached to mRNA), low-efficiency (fewer ribosomes attached) which also included mitochondrial polysomes, and the non-translating fractions. miRNAs can also associate with mRNAs that are being translated, thereby reducing the efficiency of translation, we examined the distribution of miRNAs across the fractions. The distribution of mRNAs, miRNAs, and proteins was examined under basal perfused conditions, at the end of 30 min of global no-flow ischemia, and after 30 min of reperfusion. Here we present the methods used to accomplish this analysis&#8212;in particular, the approach to optimization of protein extraction from the sucrose gradient, as this has not been described before&#8212;and provide some representative results.

Here we present a protocol to perform polysome profiling on the isolated perfused mouse heart. We describe methods for heart perfusion, polysome profiling, and analysis of the polysome fractions with respect to mRNAs, miRNAs, and the polysome proteome.

Studies in dynamic changes in protein translation require specialized methods. Here we examined changes in newly-synthesized proteins in response to ...

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.

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 ...

Biology

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, ...

Extraction of Non-Protein Amino Acids from Cyanobacteria for Liquid Chromatography-Tandem Mass Spectrometry Analysis

Non-protein amino acids (NPAAs) are a large class of amino acids (AAs) that are not genetically encoded for translation into proteins. The analysis of NPAAs can provide crucial information about cellular uptake and/or function, metabolic pathways, and potential toxicity. &#946;-methylamino-L-alanine (BMAA) is a neurotoxic NPAA produced by various algae species and is associated with an increased risk for neurodegenerative diseases, which has led to significant research interest. There are numerous ways to extract AAs for analysis, with liquid chromatography-tandem mass spectrometry being the most common, requiring protein precipitation followed by acid hydrolysis of the protein pellet. Studies on the presence of BMAA in algal species provide contradictory results, with the use of unvalidated sample preparation/extraction and analysis a primary cause. Like most NPAAs, protein precipitation in 10% aqueous TCA and hydrolysis with fuming HCl is the most appropriate form of extraction for BMAA and its isomers aminoethylglycine (AEG) and 2,4-diaminobutyric acid (2,4-DAB). The present protocol describes the steps in a validated NPAA extraction method commonly used in research and teaching laboratories.

The present protocol describes the extraction of non-protein amino acids from biological matrices via trichloroacetic acid (TCA) protein precipitation and acid hydrolysis before analysis using liquid chromatography-tandem mass spectrometry.

Non-protein amino acids (NPAAs) are a large class of amino acids (AAs) that are not genetically encoded for translation into proteins. The analysis of ...

A Streamlined Approach for Mass Spectrometry-Based Proteomics Using Selected Tissue Regions

Mass spectrometry (MS)-based proteomics enables comprehensive proteome analysis across a wide range of biological samples, including cells, tissues, and body fluids. Formalin-fixed, paraffin-embedded (FFPE) tissue sections, commonly used for long-term archiving, have emerged as valuable resources for proteomic studies. Beyond their storage benefits, researchers can isolate regions of interest (ROIs) from normal tissue regions through collaborative efforts with pathologists. Despite this potential, a streamlined approach for proteomic experiments encompassing ROI isolation, proteomic sample preparation, and MS analysis remains lacking. In this protocol, an integrated workflow that combines macrodissection of ROIs, suspension trapping-based sample preparation, and high-throughput MS analysis is presented. Through this approach, the ROIs of patients' FFPE tissues, consisting of benign serous cystic neoplasms (SCN) and precancerous intraductal papillary mucinous neoplasms (IPMN) diagnosed by pathologists, were macrodissected, collected, and analyzed, resulting in high proteome coverage. Furthermore, molecular differences between the two distinct pancreatic cystic neoplasms were successfully identified, thus demonstrating the applicability of this approach for advancing proteomic research with FFPE tissues.

Here, we establish a mass spectrometry-based proteomic method using isolated regions of interest in formalin-fixed, paraffin-embedded tissue sections. This protocol is used to analyze proteome from specific tissue areas in archived formalin-fixed, paraffin-embedded tissue sections.

Mass spectrometry (MS)-based proteomics enables comprehensive proteome analysis across a wide range of biological samples, including cells, tissues, and ...

A Protocol for the Identification of Protein-protein Interactions Based on 15N Metabolic Labeling, Immunoprecipitation, Quantitative Mass Spectrometry and Affinity Modulation

Protein-protein interactions are fundamental for many biological processes in the cell. Therefore, their characterization plays an important role in current research and a plethora of methods for their investigation is available1. Protein-protein interactions often are highly dynamic and may depend on subcellular localization, post-translational modifications and the local protein environment2. Therefore, they should be investigated in their natural environment, for which co-immunoprecipitation approaches are the method of choice3. Co-precipitated interaction partners are identified either by immunoblotting in a targeted approach, or by mass spectrometry (LC-MS/MS) in an untargeted way. The latter strategy often is adversely affected by a large number of false positive discoveries, mainly derived from the high sensitivity of modern mass spectrometers that confidently detect traces of unspecifically precipitating proteins. A recent approach to overcome this problem is based on the idea that reduced amounts of specific interaction partners will co-precipitate with a given target protein whose cellular concentration is reduced by RNAi, while the amounts of unspecifically precipitating proteins should be unaffected. This approach, termed QUICK for QUantitative Immunoprecipitation Combined with Knockdown4, employs Stable Isotope Labeling of Amino acids in Cell culture (SILAC)5 and MS to quantify the amounts of proteins immunoprecipitated from wild-type and knock-down strains. Proteins found in a 1:1 ratio can be considered as contaminants, those enriched in precipitates from the wild type as specific interaction partners of the target protein. Although innovative, QUICK bears some limitations: first, SILAC is cost-intensive and limited to organisms that ideally are auxotrophic for arginine and/or lysine. Moreover, when heavy arginine is fed, arginine-to-proline interconversion results in additional mass shifts for each proline in a peptide and slightly dilutes heavy with light arginine, which makes quantification more tedious and less accurate5,6. Second, QUICK requires that antibodies are titrated such that they do not become saturated with target protein in extracts from knock-down mutants.
Here we introduce a modified QUICK protocol which overcomes the abovementioned limitations of QUICK by replacing SILAC for 15N metabolic labeling and by replacing RNAi-mediated knock-down for affinity modulation of protein-protein interactions. We demonstrate the applicability of this protocol using the unicellular green alga Chlamydomonas reinhardtii as model organism and the chloroplast HSP70B chaperone as target protein7 (Figure 1). HSP70s are known to interact with specific co-chaperones and substrates only in the ADP state8. We exploit this property as a means to verify the specific interaction of HSP70B with its nucleotide exchange factor CGE19.

A Protocol for the Identification of Protein-protein Interactions Based on 15N Metabolic Labeling, Immunoprecipitation, Quantitative Mass Spectrometry and Affinity Modulation

We present a variation of the QUICK (QUantitative Immunoprecipitation Combined with Knockdown) approach that was introduced previously to distinguish between true and false protein-protein interactions. Our approach is based on 15N metabolic labeling, the modulation of affinities of protein-protein interactions by the presence/absence of ATP, immunoprecipitation, and quantitative mass spectrometry.

Protein-protein interactions are fundamental for many biological processes in the cell. Therefore, their characterization plays an important role in ...

Combining Chemical Cross-linking and Mass Spectrometry of Intact Protein Complexes to Study the Architecture of Multi-subunit Protein Assemblies

Proteins interact with their ligands to form active and dynamic assemblies which carry out various cellular functions. Elucidating these interactions is therefore fundamental for the understanding of cellular processes. However, many protein complexes are dynamic assemblies and are not accessible by conventional structural techniques. Mass spectrometry contributes to the structural investigation of these assemblies, and particularly the combination of various mass spectrometric techniques delivers valuable insights into their structural arrangement.
In this article, we describe the application and combination of two complementary mass spectrometric techniques, namely chemical cross-linking coupled with mass spectrometry and native mass spectrometry. Chemical cross-linking involves the covalent linkage of amino acids in close proximity by using chemical reagents. After digestion with proteases, cross-linked di-peptides are identified by mass spectrometry and protein interactions sites are uncovered. Native mass spectrometry on the other hand is the analysis of intact protein assemblies in the gas phase of a mass spectrometer. It reveals protein stoichiometries as well as protein and ligand interactions. Both techniques therefore deliver complementary information on the structure of protein-ligand assemblies and their combination proved powerful in previous studies.

The architecture of protein complexes is essential for their function. Combining various mass spectrometric techniques proved powerful to study their assembly. We provide protocols for chemical cross-linking and native mass spectrometry and show how these complementary techniques help to elucidate the architecture of multi-subunit protein assemblies.

Proteins interact with their ligands to form active and dynamic assemblies which carry out various cellular functions. Elucidating these interactions is ...

Organic Solvent-Based Protein Precipitation for Robust Proteome Purification Ahead of Mass Spectrometry

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