Optimized Protocol for the Extraction of Proteins from the Human Mitral Valve

Summary

The protein composition of the human mitral valve is still partially unknown, because its analysis is complicated by low cellularity and therefore by low protein biosynthesis. This work provides a protocol to efficiently extract protein for the analysis of the mitral valve proteome.

Abstract

Analysis of the cellular proteome can help to elucidate the molecular mechanisms underlying diseases due to the development of technologies that permit the large-scale identification and quantification of the proteins present in complex biological systems.The knowledge gained from a proteomic approach can potentially lead to a better understanding of the pathogenic mechanisms underlying diseases, allowing for the identification of novel diagnostic and prognostic disease markers, and, hopefully, of therapeutic targets. However, the cardiac mitral valve represents a very challenging sample for proteomic analysis because of the low cellularity in proteoglycan and collagen-enriched extracellular matrix. This makes it challenging to extract proteins for a global proteomic analysis. This work describes a protocol that is compatible with subsequent protein analysis, such as quantitative proteomics and immunoblotting. This can allow for the correlation of data concerning protein expression with data on quantitative mRNA expression and non-quantitative immunohistochemical analysis. Indeed, these approaches, when performed together, will lead to a more comprehensive understanding of the molecular mechanisms underlying diseases, from mRNA to post-translational protein modification. Thus, this method can be relevant to researchers interested in the study of cardiac valve physiopathology.

Introduction

Recent evidence has altered the understanding of the roles of the many regulatory mechanisms that occur after mRNA synthesis. Indeed, translational, post-transcriptional, and proteolytic processes can regulate protein abundance and function. The dogma – which says that mRNA concentrations are proxies to those of the corresponding proteins, assuming that transcript levels are the main determinant of protein abundance – has been partially revised.Indeed, transcript levels only partially predict protein abundance, suggesting that post-transcriptional events occur to regulate the proteins within cells^1,2.

Furthermore, proteins ultimately dictate the function of the cell and therefore dictate its phenotype, which can undergo dynamic changes in response to autocrine, paracrine, and endocrine factors; blood-borne mediators; temperature; drug treatment; and disease development. Thus, an expression analysis focused on the protein level is useful to characterize the proteome and to unravel the critical changes that occur to it as part of disease pathogenesis³.

Therefore, the opportunities that proteomics present to clarify health and disease conditions are formidable, despite the existing technological challenges. The particularly promising areas of research to which proteomics can contributeinclude: the identification of altered protein expression at any level (i.e., whole cells or tissue, subcellular compartments, and biological fluids); the identification, verification, and validation of novel biomarkers useful for the diagnosis and prognosis of disease; and, hopefully, the identification of new protein targets that can be used for therapeutic purposes, as well as for the assessment of drug efficacy and toxicity⁴.

Capturing the complexity of the proteome represents a technological challenge. The current proteomic tools offer the opportunity to perform large-scale, high-throughput analysis for the identification, quantification, and validation of altered protein levels. In addition, the introduction of fractionation and enrichment techniques, aimed at avoiding the interference caused by the most abundant proteins, has also improved protein identification by including the least abundant proteins. Finally, proteomics has been complemented by the analysis of post-translational modifications, which progressively emerge as important modulators of protein function.

However, the sample preparation and protein recovery in the biological specimens under analysis still remain the limiting steps in the proteomic workflow and increase the potential for possible pitfalls⁵. Indeed,in most of the molecular biology techniques that must be optimized, the first steps are tissue homogenization and cell lysis, especially during the analysis of low-abundance proteins for which amplification methods do not exist. In addition, the chemical nature of proteins can influence their own recovery. For example, the analysis of highly hydrophobic proteins is very challenging, because they easily precipitate during isoelectric focusing, while trans-membrane proteins are almost insoluble (reviewed in Reference 5). Furthermore, the tissue composition variability creates a significant barrier to developing a universal extraction method. Finally, because almost all of the clinical specimens are of limited quantity, it is essential to enable protein preparation with maximal recovery and reproducibility from minimal sample amounts⁶.

This work describes an optimized protocol for protein extraction from the normal human cardiac mitral valve, which represents a very challenging sample for proteomic analysis. The normal mitral valve is a complex structure lying between the left atrium and the left ventricle of the heart (Figure 1). It plays an important role in the control of blood flow from the atrium to the ventricle, preventing backflow and ensuring the proper level of oxygen supply to the whole body, thus maintaining an adequate cardiac output. However, it is often considered to be an "inactive" tissue, with a low cellularity and few components, mainly in the extracellular matrix. This is because, in normal conditions, the resident valvular interstitial cells (VICs) present a quiescent phenotype with a low protein biosynthesis rate⁷.

However, it has been demonstrated that, in a pathological state, the number of VICs in the spongiosa increases and their protein synthesis is activated, together with other functional and phenotypical changes⁸. Therefore, it is not surprising that the minimal data available in the literature focus on the analysis of pathological mitral valves^9,10, in which the increased number of activated VICs might explain the relatively high number of identified proteins.

In conclusion, the present protocol may serve to develop the understanding of the pathogenic mechanisms responsible for mitral valve diseases through the study of mitral valve protein components. Indeed, a greater understanding of the underlying pathological processes could help to improve the clinical management of valve diseases, whose current indications for intervention are largely predicated on hemodynamic considerations.

Protocol

In this protocol, the human hearts are collected during multiorgan explantation (cold ischemia time of 4-12 h, mean 6 ± 2 h) from multi-organ donors excluded from organ transplantation for technical or functional reasons, despite normal echocardiographic parameters. They are sent to the Cardiovascular Tissue Bank of Milan, Monzino Cardiologic Center (Milan, Italy) for the banking of the aortic and pulmonary valves. The mitral posterior leaflets are not used for clinical purposes, so they are collected during the aortic and pulmonary valve isolation after informed consent is obtained from the donors' relatives. The tissue for transplantation and research is collected only after parental consent; on the consent sheet, they authorize (or not) the use of the cardiac tissue for research only if it is not suitable for human clinical use (i.e., microbiological, functional, and serological problems), following the guidelines of the ethics committee of Monzino Cardiologic Center.

1. Mitral valve preparation

1. Harvest the human mitral valve as soon as possible after organ explantation (cold ischemia time of 4-12 h).
2. In a clean room, remove the heart from the transport bag containing a cold (4 °C) solution (i.e., saline solution or balanced medium Eurocollins or Wisconsin). Put it into a bucket and place it in a biosafety cabinet (biohazard vertical air flow, class A, Good Manufacturing Practices (GMP) classification) to proceed with the valve preparation.
3. Place the heart on a sterile disposable drape in the cabinet. Using a sterile disposable scalpel, cut the heart completely, perpendicularly to its major axis, on the level of the left and right ventricles, about 4 cm away from the apex.
4. Move the ascending aorta and pulmonary artery to display the left atrial roof.
5. With sterile autoclavable forceps and picks, cut around the left auricle on the left atrial roof, making the mitral valve visible and allowing for the great mitral leaflet (anterior) and the small mitral leaflet (posterior) to be identified.
   NOTE: Antero-lateral and the posterior medial commissures define the border of the anterior leaflet and the posterior area.
6. Using sterile autoclavable scissors and non-traumatic forceps, dissect the left atrium and the ventricle wall thickness around the circumference of the whole mitral valve.
7. Identify the mitro-aortic valve continuity.
   NOTE: The left ventricle contains the whole mitral valve and chords.
8. Separate the anterior mitral valve leaflet from the posterior mitral valve leaflet, cutting the posterior leaflet along the insertion with the ventricle (commissure).
9. Wash the posterior leaflet in the saline solution. Cut the leaflet into small pieces (<1 cm²) and individually wrap them in aluminum foil. Snap-freeze them with liquid nitrogen.
   Caution: Follow organizational safety procedures when using liquid nitrogen.
   1. Sanitize the table of the cabinet with a 70% isopropyl alcohol solution and a 6% hydrogen peroxide solution at the end of the procedure.

2. Protein extraction

1. Use forceps to pick up the sample stored in liquid nitrogen and immediately place it on dry ice while still wrapped in the aluminum foil. Do not leave the sample to thaw during any transfers.
2. Prior to grinding, chill the porcelain/zirconium mortar and pestles of a grinder system (e.g., CryoGrinder), together with the sample, by putting them in a Dewar flask containing liquid nitrogen (~500 mL).
   Caution: Follow organizational safety procedures when using liquid nitrogen.
3. Put the mortar and pestles in a polystyrene box containing dry ice. Remove the sample from the aluminum foil and put it into the mortar.
4. Grind the sample with the big pestle against the mortar 15-20 times, using the screwdriver to rotate the pestle. Mix the sample with the tip of a pre-chilled spatula during the grinding process.
   1. Repeat with the small pestle.
5. Transfer the ground sample to a previously weighed tube (e.g., 15 mL centrifuge tube) by inverting the tube, placing it over the mortar, and inverting them together to move the sample to the tube. Use a pre-chilled spatula to recover all material from the mortar.
   1. Keep the tube with the sample on dry ice to avoid sample thawing during the transfer.
6. Calculate the net weight of the sample.
7. Clean the mortar and the pestles after each sample and decontaminate them by autoclaving or heating them at 200 °C for 2 h.
8. Transfer the powdered sample from the centrifuge tube to the glass tube of a homogenizer by inversion.
9. Add filtered urea buffer (8 M urea, 2 M thiourea, 4% w/v CHAPS, 20 mM Tris, and 55 mM dithiotreitol) to the glass tube, 200 µL of urea buffer for every 10 mg of powdered tissue.
   NOTE: Residual powdered sample left in the centrifuge tube can be recovered using part of the calculated volume of urea buffer.
10. Homogenize the sample using a stirrer equipped with a borosilicate glass mortar and a polytetrafluoroethylene (PTFE) pestle. Slowly press the pestle onto the sample with a twisting motion (1,500 rpm) 10 times.
11. Recover the supernatant and transfer it to a clean 1.7 mL centrifuge tube. Again extract the remaining sample with fresh urea buffer, adding half of the volume used during the first extraction.
12. Repeat step 2.10.
13. Recover the supernatant and combine it with the supernatant from step 2.11. Place the combined supernatant on a tube rotator for 30 min.
14. Centrifuge the tube for 30 min at 13,000 x g and 4 °C.
15. Recover the supernatant and measure the protein concentration using the Bradford protein assay, per the manufacturer's instructions. Store the sample at -80 °C until use.

Results

The extraction and dissolution of proteins in the urea buffer is directly compatible with proteomic methods based on isoelectrofocusing (two-dimensional electrophoresis (2-DE)¹¹ and liquid-phase isoelectric focusing (IEF)¹²) and with immunoblotting after dilution in Laemmli buffer¹³ containing a protease inhibitor cocktail¹⁴.

For gel-...

Discussion

One critical step of this protocol is the use of liquid nitrogen to freeze the sample and to chill the grinder system. The use of liquid nitrogen prevents biological degradation and allows for efficient powdering, but it requires specific training for safe handling.

In this protocol features a grinder system for sample grinding because small samples are difficult to recover from standard mortar and pestles. In this case, small samples spread as a fine powder over the mortar surface, rendering ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Saline solution			0.9 % NaCl
Eurocollins A	SALF	30874046	Balanced organ's transport medium. Combine 400 mL of Eurocollins A with 100 mL Eurocollins B to obtain balanced medium Eurocollins
Eurocollins B	SALF	30874022	Balanced organ's transport medium. Combine 400 mL of Eurocollins A with 100 mL Eurocollins B to obtain balanced medium Eurocollins
Wisconsin	Bridge life	RM/N 4081	Balanced organ's transport medium
Biohazard vertical flow air	Burdinola		Class A GMP classification
Dewar Flask	Thermo Scientific	Nalgene 4150-1000
Cryogrinder system	OPS diagnostics	CG 08-01	Grinder system containing mortars, pestles and screwdriver
Stainless steel forceps
Stainless steel spatula
Disposable sterile scalpel	Medisafe	MS-10
Stainless steel scissors			Autoclavable
Stainless steel picks			Autoclavable
Disposable sterile drap	Mon&Tex	3.307.08
Sterilizing solution with isopropyl alcohol			70% isopropyl alcohol
Sterilizing solution with hydrogen peroxide			6% hydrogen peroxide
Micropipette, 1 mL, with tips
15 mL centrifuge tubes	VWR international	9278
1.7 mL centrifuge tubes	VWR international	PIER90410
Urea buffer			8 M urea, 2 M thiourea, 4 % w/v CHAPS, 20 mM Trizma, 55 mM Dithiotreitol
Urea	Sigma aldrich	U6504-1KG	To be used for Urea buffer
Thiourea	Sigma aldrich	T8656	To be used for Urea buffer
CHAPS	Sigma aldrich	C3023-5GR	To be used for Urea buffer
Dithiotreitol	Sigma aldrich	D0632-5G	To be used for Urea buffer
Syringe 50 mL	PIC		To be used to filter Urea buffer
0.22 µm filter	Millipore	SLGP033RB	To be used to filter Urea buffer
PFTE Pestle, 2 mL	Kartell	6302	Part of Potter-Elvehjem homogenizer
Borosilicate glass mortar	Kartell	6102	Part of Potter-Elvehjem homogenizer
Stirrer	VELP scientifica	Stirrer DLH	To be used for homogenization by Potter-Elvehjem
Bradford Protein assay	Bio-Rad laboratories	5000006
Tube rotator	Pbi International	F205
Liquid nitrogen
Aluminum foil
Ice
Polystyrene box
Dry ice
Centrifuge			For centrifugation of 1.7 mL centrifuge tubes at 13,000 x g
Freezer -80°C
Precision balance
Autoclave			For sterilization
Cryogenic gloves for liquid nitrogen
Gloves
Professional forced ventilation and natural air convection oven			For sterilization
Protease inhibitor cocktail	Sigma aldrich	P8340-5ML	100X solution
ProteoExtract Protein Precipitation Kit	Calbiochem	539180
RapiGest	Waters	186001861
Cytoscape	www.cytoscape.org	version 2.7	Software platform for Gene Ontology analysis
BiNGO	http://apps.cytoscape.org/apps/bingo	version 3.0.3	Plugin for Gene ontology analysis
AlphaB Crystallin/CRYAB Antibody	Novus Biologicals	NBP1-97494	Mouse monoclonal antibody against CryAB
Septin-11 Antibody	Novus Biologicals	NBP1-83824	Rabbit polyclonal antibody against septin-11
FHL1 Antibody	Novus Biologicals	NBP-188745	Rabbit polyclonal antibody against FHL-1
Dermatopontin Antibody	Novus Biologicals	NB110-68135	Rabbit polyclonal antibody against dermatopontin
Goat Anti mouse IgG HRP	Sigma aldrich	A4416-0.5ML	Secondary antibody for immunoblotting
Goat Anti rabbit IgG HRP	Bio-Rad laboratories	170-5046	Secondary antibody for immunoblotting