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Keywords:

  • Animal proteomics;
  • Biomineralization;
  • Bleaching treatment;
  • Calcifying extracellular matrix;
  • Protein identification;
  • Sample preparation

Abstract

  1. Top of page
  2. Abstract
  3. 1 Introduction
  4. 2 Contamination of extracellular calcifying matrices extracted from biomineral structure by cellular components
  5. 3 Why it is crucial to avoid cellular component contamination and generate specific protein lists
  6. 4 Our recommendations on how to analyze proteins from biomineralized structures by proteomics
  7. 5 Conclusion
  8. Acknowledgment
  9. 6 References

In a recent editorial (Proc. Natl. Acad. Sci., 2013 110, E2144–E2146) and elsewhere, questions have been raised regarding the experimental practices in relation to the proteomic analysis of organic matrices associated to the biomineralized CaCO3 skeletons of metazoans such as molluscan shells and coral skeletons. Indeed, although the use of new high sensitivity MS technology potentially allows to identify a greater number of proteins, it is also equally (or even more) sensitive to contamination of residual proteins from soft tissues, which are in close contact with the biomineral. Based on our own past and present experimental know-how—observations that are reproducible and coherent with the current understanding of extracellular biomineralization processes—we are convinced that a careful and appropriate cleaning of biominerals prior to any analysis is crucial for accurate proteomic investigations and subsequent pertinent interpretation of the results. Our goal is to alert the scientific community about the associated bias that definitely should be avoided, and to provide critical recommendations on sample preparation and experimental design, in order to better take advantage of the aptness of proteomic approaches aiming at improving our understanding of the molecular mechanisms in biomineralization.

Abbreviations
ECM

extracellular matrix

NaOCl

sodium hypochlorite

RLCD

repeat low complexity domain

SOMP

skeleton organic matrix protein

1 Introduction

  1. Top of page
  2. Abstract
  3. 1 Introduction
  4. 2 Contamination of extracellular calcifying matrices extracted from biomineral structure by cellular components
  5. 3 Why it is crucial to avoid cellular component contamination and generate specific protein lists
  6. 4 Our recommendations on how to analyze proteins from biomineralized structures by proteomics
  7. 5 Conclusion
  8. Acknowledgment
  9. 6 References

Most of metazoan skeletons are produced extracellularly via the secretion of precursor ions, together with an acellular organic matrix that remains embedded within the mature biomineral structures once deposited. This extracellular matrix (ECM) comprises amalgamates of proteins, glycoproteins, polysaccharides, and lipids, with the proteinaceous fraction being the dominant moiety. Although the ECM represents only a small fraction of the biomineral weight (between 0.1 and 5% w/w), it is thought to exquisitely regulate the mineral deposition, and consequently, to play a central role in the whole biomineralization process [1].

Since the 1990s, ECM proteins extracted from calcified ske-letons of few nonvertebrate metazoan models have been gradually characterized by “one-by-one” approaches (for review on echinoderms or mollusks see [2-4]). However, this strategy did not give a complete picture of these ECM protein repertoires. More recently, thanks to advances in high-throughput genomic and transcriptomic sequencing of an increasing number of nonmodel organisms, proteomic analyses of the so-called skeleton organic matrix proteins (SOMPs) extracted after dissolution of the mineral phase, combined with the interrogation of nucleic acid datasets has resulted in the description of numerous novel proteins from various nonvertebrate metazoan species [5-14].

Correspondence concerning this and other Viewpoint articles can be accessed on the journals' home page at: http://viewpoint.proteomics-journal.de

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The combination of global SOMP MS-based proteomics and transcriptomics/genomics performed by us [6, 7, 10, 12, 13] has led to the identification of about 40–60 ECM-specific proteins (depending on the biological model), exhibiting unique primary structures with signal peptide, transmembrane and repeat low-complexity domains, enzyme and/or ECM signatures. In addition, a specific expression of their transcripts can be measured in skeleton-secreting tissues or by immunolocalization of the translated proteins [10, 12], constituting a strong experimental evidence of their involvement in the biomineralization process. Surprisingly, few other works have published much larger lists of biomineral ECM-associated proteins that were identified employing similar approaches (up to 200–300 proteins per model depending on the taxa [9, 11]). But contrary to our findings, the latter lists contain, in addition to ECM-specific proteins, numerous intracellular proteins. In our experiments, these obvious cell constituent proteins are not observed when biomineral structures are adequately cleaned prior to ECM extraction. Hence, we assert that these proteins should be considered as contaminants, and not assigned as true SOMPs without further investigation [15].

2 Contamination of extracellular calcifying matrices extracted from biomineral structure by cellular components

  1. Top of page
  2. Abstract
  3. 1 Introduction
  4. 2 Contamination of extracellular calcifying matrices extracted from biomineral structure by cellular components
  5. 3 Why it is crucial to avoid cellular component contamination and generate specific protein lists
  6. 4 Our recommendations on how to analyze proteins from biomineralized structures by proteomics
  7. 5 Conclusion
  8. Acknowledgment
  9. 6 References

During the editorial process of one of our previous manuscripts on the proteomic investigation of the calcified shell layers from the gastropod Haliotis asinina [6], an anonymous reviewer asked us to justify why we identified only ECM-specific proteins and no intracellular ones, arguing that biomineralization does not take place in a clean room, and that intracellular proteins are often part of the list of ECM's proteins, as exemplified by the work pertaining to the calcified structures of the sea urchin Strongylocentrotus purpuratus [5]. Although, intracellular proteins (such as actins and tubulins) can be sometimes detected within the extractable organic components associated to mineralized tissues [14], we assume that their occurrence results from contamination by cell constituting proteins of cellular remains [13, 15], and are not embedded or strongly associated to the mineral phase. Indeed, other proteomic analyses on the organic matrices extracted from cleaned otolith (from fish) led to the identification of only few specific proteins (Table 1), all ECM-related, that appear to be directly involved in the formation of these inner ear calcified structures [16].

Table 1. Summary of the main MS-based proteomic approaches applied in the recent years (2005–2013) to mineralized structures of calcium carbonate from metazoan origin
 SpeciesCalcifiedKey cleaningDemineralizationOrganicExtractionLC-MS/MS/DataIdentifiedProtein
  tissuesteps fractionsprocedureinterrogationsourceproteinslocalization
  1. Accession numbers of identified proteins were collected from the literature and corresponding sequence data resources. The subcellular location was predicted by means of bioinformatics tools (http://www.cbs.dtu.dk/services) according to the protocol described in [17]. In brief, protein sequences were analyzed with TargetP, TMHMM for transmembrane domains, SignalP for the presence of peptide signals, GPI for the presence of GPI anchors and SecretomeP for nonclassical secretion. Complete protein sequences without potential secretory and/or membrane signatures were considered intracellular without further characterization of their predicted location inside the cell. As for protein fragments, localization was predicted by gene ontology when available or considered unknown otherwise.

 Cnidaria         
 Acropora millepora [13]Skeleton
  1. Washed fragments in NaOCl 5% v/v, 72 h.
  2. Powder (<200 μm) in NaOCl 1% v/v, 5 h.
Acetic acid 10% v/v overnight at 4°C until pH 4ASM
  • Centrifugation
  • Ultrafiltration
  • Dialysis
LTQ–FT/MASCOT
  • NCBI
  • nucleotides (101 380)
  • EST (15 389)
36
  • ECM—15
  • ECM/ Membrane—13
  • Membrane—3
  • Intracellular—0
  • Unknown—5
     AIM
  • 6 × Centrifugation with milliQ water
    
 Acropora millepora [13, 15]Skeleton
  1. Washed fragments in NaOCl 5% v/v, 72 h.
Acetic acid 10% v/v overnight at 4°C until pH 4ASM
  • Centrifugation
  • Ultrafiltration
  • Dialysis
LTQ–FT/MASCOT
  • NCBI
  • nucleotides (101 380)
  • EST (15 389)
52
  • ECM—12
  • ECM/ Membrane—7
  • Membrane—5
  • Intracellular—14
  • Unknown—14
 Stylophora pistillata [14]Skeleton
  1. Washed fragments in NaOCl 3% wt/v, 4 h.
  2. Powder (<150 μm) second bleaching.
1 N HCl at room temperature pH 7
  • ASM
  • AIM
  • Centrifugation
  • Acetone 90%
  • Centrifugation
LTQ–FT/X! TandemDraft genome36
  • ECM—7
  • ECM/Membrane —0
  • Membrane—8
  • Intracellular—7
  • Unknown—14
 Echinodermata         
 Strongylocentrotus purpuratus [5]SpiculesIsolation of spicules followed by centrifugation interspersed by successive resuspensions in:Acetic acid 50% v/v for 5 h at 4°CASM
  • Dialysis
LTQ–FT/ MaxQuantPredicted annotated protein models (Glean3)231
  • ECM –72
  • ECM/Membrane—40
  • Membrane—51
  • Intracellular—66
  • Unknown—2
   
  1. NaOCl 4.5% v/v for 1–2 min.
  2. CaCO3-saturated water.
  3. Ethanol, then acetone (100%).
       
  Test and spine
  1. Tests cut into 2 halves and washed.
  2. 3 × 200 mL NaOCl (6–14% active chlorine), 10 min.
Acetic acid 50% v/v overnight at 4°CASMASM
  • Dialysis
LTQ-FT/ MASCOTPredicted annotated protein models (Glean3)110
  • ECM—35
  • ECM/Membrane—23
  • Membrane—20
  • Intracellular—28
  • Unknown—4
  Tooth
  1. 4 × 200 mL NaOCl (6–14% active chlorine), 1 h, with changes after 15 min with a 2-min sonication interval after every change.
  2. Reduced to powder and washed again as in 1.
Acetic acid 50% v/v overnight at 4°CASM
  • Dialysis
LTQ-FT/ MASCOTPredicted annotated protein models (Glean3)138
  • ECM—49
  • ECM/ Membrane—24
  • Membrane—40
  • Intracellular—21
  • Unknown—4
 Mollusca         
 Pinctada margaritifera and Pinctada maxima [10]
  • Shell:
  • Nacre
  • Prisms
  1. Intact shells in NaOCl 1% v/v for 24 h.
  2. Separated shell layers thoroughly rinsed with water, crushed into ∼1-mm2 fragments and subsequently into fine powder (>200 μm).
Acetic acid 5% v/v overnight at 4°C until pH 4.2ASM
  • Centrifugation
  • Ultrafiltration
  • Dialysis
Q-TOF/ MASCOT and Protein-Pilot
  • NCBI
  • EST (76 790) – P. margaritifera
  • EST + nucleotide (7272) – P. maxima
80
  • ECM—51
  • ECM/ Membrane—4
  • Membrane—6
  • Intracellular—0
  • Unknown—19
     AIM
  • 6 × Centrifugation with milliQ water
    
 Haliotis asinina [6]
  • Shell:
  • Nacre
  • Prisms
  1. Intact shells in NaOCl 1% v/v for 24 h.
  2. Separated shell layers thoroughly rinsed with water, crushed into ∼1-mm2 fragments and subsequently into fine powder (>200 μm).
Acetic acid 5% v/v overnight at 4°C until pH 4.2ASMAIM
  • Centrifugation
  • Ultrafiltration
  • Dialysis
  • 6 × Centrifugation with milliQ water
Q-TOF/ MASCOT
  • NCBI
  • Nucleotides + EST (9.167)
14
  • ECM—11
  • ECM/ Membrane—0
  • TM—0
  • Intracellular—0
  • Unknown—3
 Crassostrea gigas [11]
  • Shell:
  • Nacre
  • Prisms
  1. Intact shells in NaOCl, 24 h.
30 mL of acetic acid solution 5% until pH 4.0, stirred overnight.Not specified
  • TCA 20%, 2h
  • Centrifugation
  • (3×) Acetone andcentrifugation
LTQ-FT/ MASCOT
  • NCBI
  • Annotated protein models (26 086)
259
  • ECM—75
  • ECM/ Membrane—10
  • Membrane—30
  • Intracellular—135
  • Unknown—9
 Lottia gigantea [9]
  • Shell:
  • Spherulitic Prismatic
  • Cross-lamellar
  1. Intact shells in NaOCl (6–14% active chlorine) for (A) 2 h at RT, (B) 2 h with ultrasound, (C) 24 h with ultrasound
Acetic acid 50% v/v overnight at 4–6°C
  • ASM
  • AIM
  • 2× Dialysis
LTQ-FT/ MaxQuant
  • genome.jgi-psf. org/
  • Annotated protein models (23 851)
311
  • ECM—141
  • ECM/ Membrane—20
  • TM—31
  • Intracellular—89
  • Unknown—30
 Lottia gigantea [12]
  • Shell:
  • Prismatic Cross-lamellar
  1. M + 2, M + 1, M and M − 1 layers were crushed into approximately 1-mm2 fragments
  2. Shell fragments in NaOCl 1% v/v, 24 h.
Acetic acid 5% v/v overnight at 4°C until pH 4.2ASM
  • Centrifugation
  • Ultrafiltration
  • Dialysis
Q-TOF/ MASCOT
  • NCBI
  • Nucleotides+EST (252 091)
  • genome.jgi-psf. org/
  • Annotated protein models (23 851)
39
  • ECM—31
  • ECM/ Membrane—3
  • Membrane—3
  • Intracellular—0
  • Unknown—2
     AIM
  • 6 × Centrifugation with milliQ water
    
 Chordata         
 Gallus gallus [18]Eggshell
  1. Isolated eggshells in 5% EDTA and then washed with water.
Acetic acid 10% v/vASM
  • Centrifugation
  • Dialysis
LTQ-FT/ MASCOT
  • EBI
  • chicken IPI protein sequence database
  • (∼ 25 772)
520
  • ECM—226
  • ECM/ Membrane—43
  • Membrane—75
  • Intracellular—140
  • Unknown—36
 Danio rerio and Oncorhynchus mykiss [16]Otolith
  1. Isolated otolith in 0,65% sodium hypochlorite, then sonicated and washed with water.
EDTA excessESM and EISM
  • Centrifugation
  • Ultrafiltration
Q-TOF/ MASCOT
  • NCBI, Ensembl
  • Fish genomes
  • (∼ 2 106 626)
8
  • ECM—7
  • ECM/ Membrane—0
  • Membrane—1
  • Intracellular—0
  • Unknown—0

To better illustrate this problem, we report on Table 1 the main recent proteomic approaches applied to study mineralized structures of CaCO3 in four metazoan phyla, describing the cleaning procedure, demineralization and extraction steps together with the corresponding database search tools used for protein identification. By collecting the number of proteins identified in each study and inferring their predicted location based on sequence properties [17] it is clear that the cleaning step may, at least, partially, strongly influence the number of hits, in particular, it can increase the number of identifications corresponding to intracellular constituents and other ubiquitous proteins. We believe however that these protein hits can be avoided by an extensive and adapted cleaning method of the biomineral structures, prior to the extraction of the organic matrix.

In addition to the information described in Table 1, we explain here the effect of the cleaning procedure typified by two examples: the first one deals with the investigation of the SOMP from freshwater gastropod Lymnaea stagnalis (Fig. 1). We demonstrate that some proteins, such as actins, tubulins, ATPases, and myosins, are intracellular contaminants from cell fragments, that remain after a “simple bleaching” treatment, but can be removed by an additional drastic cleaning of the biomineral fine powder with concentrated sodium hypochlorite (NaOCl) (10%, 5 h) in addition to the umbilicus removal. The second example refers to the SOMP analysis of the stony coral Acropora millepora [13, 15], for which—similarly to the first example—two bleaching treatments were required to remove cellular contaminants. We would like to insist here on the fact that the publication of SOMP lists containing such contaminants is misleading and detrimental for our understanding of biocalcification mechanisms and to the elaboration of molecular models, since there is currently no evidence that intracellular proteins—no matter their subcellular localization—interact directly with the growing biomineral. Moreover, because it blurs the picture of the diversity of SOMPs, and the comprehension of ECM functions in biomineralization processes, this problem aims at being carefully appreciated.

image

Figure 1. Removal of organic contaminants from biomineralized shell structures of Lymnaea stagnalis. Comparison of the proteins identified by proteomics on the skeletal organic matrix under two different conditions. “Simple bleaching” consisted of treating the skeletal fragments with NaOCl solution once (5% v/v, 24 h), while in “extended bleaching” the “simple bleaching” was followed by complete removal of the entire umbilicus shell portion and a subsequent treatment with NaOCl solution (10% v/v, 5 h) on the skeletal sieved powder (<200 μm). The acetic acid-soluble and -insoluble matrices were digested with trypsin and were injected into a Q-Star XL nano-electrospray quadrupole/TOF tandem mass spectrometer then protein identifications were performed with the Mascot search engine against specific transcriptomic datasets.

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3 Why it is crucial to avoid cellular component contamination and generate specific protein lists

  1. Top of page
  2. Abstract
  3. 1 Introduction
  4. 2 Contamination of extracellular calcifying matrices extracted from biomineral structure by cellular components
  5. 3 Why it is crucial to avoid cellular component contamination and generate specific protein lists
  6. 4 Our recommendations on how to analyze proteins from biomineralized structures by proteomics
  7. 5 Conclusion
  8. Acknowledgment
  9. 6 References

One major point for understanding organic matrix-mediated biomineralization processes is to identify, for a given biological model, all key proteins (the “minimal toolbox”) required for mineralization and their respective functions. The proteomic investigations of SOMPs constitute a valuable approach to reach this goal, and gradually provide fundamental insights on the molecular basis of biomineralization [6, 10, 12]. In practice, only the investigation of genuine SOMPs—directly associated with the mineral phase formation—such as the example of caspartin, an intracrystalline protein from Pinna nobilis calcitic prisms, can contribute to important fundamental discoveries [19]. Moreover, understanding how these proteins have evolved, by a comparative approaches on different models, is also likely to provide new insights into the events that supported the organism evolution [12]. In this specific context, the presence of contaminating cytoskeletal proteins (e.g. actins, tubulin, myosin, …) and more generally of intracellular components (e.g. histones, ribosomal proteins, …) is particularly misleading. The case of actin, one of the most important cytoskeletal proteins, is representative of the problem, since it was included in the so-called SOMP list from different species [14, 20], suggesting homologous mechanisms associated with biomineralization processes without further substantial evidence.

For the proteins presenting obvious transmembrane domains, it appears that some of them may also participate in the biomineralization process (Table 1), by means of the domains that are targeted outside the cell. These extracellular regions may potentially act in the mineralizing space being subsequently cleaved by proteases and occluded in the growing biomineral. We present evidences supporting this hypothesis in the works of Ramos-Silva et al. [13, 15]. The same scenario is conveniently suggested by Mann et al. [5], when performing a global proteomics analysis on the spicule organic matrix of the sea urchin.

Until now, we have considered that the data about ECM contamination did not deserve to be included in our publications that concerns the identification of biomineral-associated proteins by proteomics [6, 7, 10, 12, 13], however the misleading interpretation of some potential contaminants described as SOMPS that are being published in an increasing number of research articles [9, 11, 14, 20, 21], urge us to inform the scientific community about this issue.

4 Our recommendations on how to analyze proteins from biomineralized structures by proteomics

  1. Top of page
  2. Abstract
  3. 1 Introduction
  4. 2 Contamination of extracellular calcifying matrices extracted from biomineral structure by cellular components
  5. 3 Why it is crucial to avoid cellular component contamination and generate specific protein lists
  6. 4 Our recommendations on how to analyze proteins from biomineralized structures by proteomics
  7. 5 Conclusion
  8. Acknowledgment
  9. 6 References

According to the most commonly accepted view, the formation of the metazoan external calcified structures results from the secretion of an acellular matrix that remains occluded within the biomineral phase once precipitated. Alternatively, some other calcified structures, such as the sclerites produced by soft corals [20], are formed, in a first step, inside intracellular vesicles, then being externalized by exocytosis. During these extracellular and/or confined intracellular processes, cellular contaminants can remain entrapped in the mineralized structures, such as the microcavities present in some mollusk shell layers (Fig. 2A and B). Besides the specimen cellular remains, another noticeable source of contamination is exogenous and is mostly represented by the microorganisms that can grow within the exoskeletal cavities, such as the various specific endolith algae of stony corals [22] or the boring groups (bacteria, sponge, or algae) that can live in some mollusk shells.

image

Figure 2. Details of two mollusk shell layers containing void areas that can entrap organic contaminants, as exemplified for the chalky layer of the Pacific oyster Crassostrea gigas (A) or the bioperforated external prismatic layer of the giant limpet Lottia gigantea (B). These structures potentially entrap endogenous (e.g. cellular debris from the mantle cells) or exogenous (e.g. bacteria or other parasitic micro-organisms) contaminant organic material.

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As a first cleaning step, we recommend to mechanically remove (by abrasion or sawing) such “potentially rich in contaminant” zones, in order to minimize contamination, or to critically appreciate the data obtained, when primarily analyzing biominerals by proteomics. Considering the microstructural diversity of bioconstructions produced by living organisms, we recommend the development of optimized cleaning methods for the analysis of all new biomaterials, based on the detailed characterization of these microstructures. A bleaching treatment in two steps may assure the quality of the proteomic data: first one for removing most of the superficial contaminants, a second one for “flushing out” the most occluded ones. For instance, we observed that for some biominerals, only a thorough incubation of skeleton fine powder (<200 μM)—and not simply of skeletal fragments—in concentrated NaOCl (10%, 5 h), before extraction, was fully efficient at removing contaminants, leaving intact the skeleton associated proteins, i.e. true SOMPs that are part of the “biomineralization toolkit” [10, 12, 13]. We recommend the users to adopt similar cleaning procedures. Rather than using sodium hydroxide or hydrogen peroxide, we recommend NaOCl, which offers the compromise between being a very effective bleaching agent without dissolving the mineral phase [23], but this should be tested according to the different biominerals investigated.

Generally, considering the diversity of CaCO3 biominerals, one should consider that there is not one universal cleaning protocol for biomineralized structures. Protocols need to be optimized by adjusting the concentration of the cleaning agent, the duration of the treatment, and the grain size of the powder, for proteomic approaches on new biominerals. One disadvantage of sample cleaning is that it can also potentially discard proteins of interest that are “weakly” bound to the mineral phase. However, our previous analysis indicates that cytoskeletal and intracellular components are the ones that are predominantly removed by NaOCl treatment [13, 15]. Comparative approaches of the sample cleaning procedures would also lead to discriminate between biomineral-associated proteins that are weakly bound around the biomineral without being occluded, and “true” SOMPs, that are specifically integrated inside the mineral phase. Beside the cleaning procedure, attention should be accorded to the subsequent analytical steps:

  1. Many known SOMPs present remarkable bias in their amino acid composition, presence of multiple repeated low-complexity domains (RLCDs), higher insolubility, lack of trypsin cleavage sites. This requires the development of novel protein digestion strategies (e.g. enzyme-, microwave-assisted digestions, or acid hydrolysis) that allow generating peptides of optimal length for MS/MS analysis [24].
  2. It is also worth to note that the specific amino acid composition of many SOMP influences the results of the peptide-spectrum matches according to chosen fragmentation technique (CID, higher energy dissociation, electron transfer dissociation), and as well as the performance of the search engines [25].
  3. Often, SOMPs present PTMs (e.g. glycosylations, phosphorylations, …) which can limit the access to the protein cleavage sites, and induce poorer ionization, resulting in lesser protein identifications. As the amount of saccharidic moieties can significantly differ according to the investigated species, chemical or enzymatic deglycosylation step of biomineral-extracted matrix may be recommended in some cases.
  4. The completeness of the database for SOMP encoding transcripts, either from genome or transcriptome assembly, is especially critical for the identification of a greater number of SOMPs. The availability of mineralizing tissue-specific transcriptome datasets represents also a great advantage. However few examples highlight that these resources can sometimes be incomplete and not always sufficient to identify all SOMPs [6, 12].

In addition to the above points, critical analysis of the characteristics of the identified proteins is required, which includes the following criteria: the specific expression of their transcript in mineralizing tissues, the presence of predicted signal peptide, and transmembrane- or ecto-domains, their sequence similarity with other biomineralization proteins, and functional characterization according to experimental evidence or, alternatively, to gene ontology determination using sequence comparison approach.

5 Conclusion

  1. Top of page
  2. Abstract
  3. 1 Introduction
  4. 2 Contamination of extracellular calcifying matrices extracted from biomineral structure by cellular components
  5. 3 Why it is crucial to avoid cellular component contamination and generate specific protein lists
  6. 4 Our recommendations on how to analyze proteins from biomineralized structures by proteomics
  7. 5 Conclusion
  8. Acknowledgment
  9. 6 References

The purpose of this viewpoint is to alert the scientific community about the proper approach for analyzing skeletal organic matrix proteins by proteomics, and to get better benefit of this approach. Indeed, incorrect interpretation of contaminants as genuine SOMPs can potentially blur the biological patterns that may emerge from the analysis, and may mask fascinating evolutionary and/or functional trends. From our viewpoint, contaminations are technically possible to limit, if not to avoid, but require scrupulous and accurate sample treatments.

Moreover, it is obvious that the use of more and more sensitive MS technologies (e.g. Orbitrap), leads to greater protein scoring but also increases the susceptibility to detect associated contaminants present even in very small amounts, then the limit between true SOMPs present in small amount, and contaminants, would be even more difficult to draw.

Acknowledgment

  1. Top of page
  2. Abstract
  3. 1 Introduction
  4. 2 Contamination of extracellular calcifying matrices extracted from biomineral structure by cellular components
  5. 3 Why it is crucial to avoid cellular component contamination and generate specific protein lists
  6. 4 Our recommendations on how to analyze proteins from biomineralized structures by proteomics
  7. 5 Conclusion
  8. Acknowledgment
  9. 6 References

The authors would like to thank Dr. Isabelle Zanella-Cléon (Institut de Biologie et de Chimie des Protéines, Lyon France) for the mass spectrometry analysis, Jérome Thomas (Université de Bourgogne, Dijon, France) for picture processing. Lymnaea stagnalis shells were provided by Dr. Daniel John Jackson from Göttingen University, Germany.

The authors have declared no conflict of interest.

6 References

  1. Top of page
  2. Abstract
  3. 1 Introduction
  4. 2 Contamination of extracellular calcifying matrices extracted from biomineral structure by cellular components
  5. 3 Why it is crucial to avoid cellular component contamination and generate specific protein lists
  6. 4 Our recommendations on how to analyze proteins from biomineralized structures by proteomics
  7. 5 Conclusion
  8. Acknowledgment
  9. 6 References
  • 1
    Baeuerlein, E., in: Baeuerlein, E. (Ed.), Handbook of Biomineralization—Biological Aspects and Structure Formation, Wiley-VCH Verlag GmbH & Co KGaA, Weinheim, Germany 2007, pp. 119.
  • 2
    Killian, C. E., Wilt, F. H., Molecular aspects of biomineralization of the echinoderm endoskeleton. Chem. Rev. 2008, 108, 44634474.
  • 3
    Marin, F., Luquet, G., Marie, B., Médakovic, D., Molluscan shell protein: primary structure, origin and evolution. Curr. Top. Dev. Biol. 2008, 80, 209276.
  • 4
    Marin, F., Marie, B., Benhamada, S., Silva, P., et al., ‘Shellome’: proteins involved in mollusk shell biomineralization – diversity, functions, in: Watabe, S., Maeyama, K., Nagasawa, H. (Eds.), Recent Advances in Pearl Research, Terrapub, Tokyo, Japan 2013, pp. 149166.
  • 5
    Mann, K., Wilt, F. H., Poustka, A. J., Proteomic analysis of sea urchin (Strongylocentrotus purpuratus) spicule matrix. Proteome Sci. 2010, 8, 33.
  • 6
    Marie, B., Marie, A., Jackson, D., Dubost, L. et al., Proteomic analysis of the organic matrix of the abalone Haliotis asinina calcified shell. Proteome Sci. 2010, 8, 54.
  • 7
    Joubert, C., Piquemal, D., Marie, B., Manchon, L. et al., Transcriptome and proteome analysis of Pinctada margaritifera calcifying mantle and shell: focus on biomineralization. BMC Genomics 2010, 11, 613.
  • 8
    Berland, S., Marie, A., Duplat, D., Milet, C. et al., Coupling proteomics and transcriptomics for the identification of novel and variant forms of mollusc shell proteins: a study with P. margaritifera. Chembiochem. 2011, 12, 950961.
  • 9
    Mann, K., Edsinger-Gonzales, E., Mann, M., In-depth proteomic analysis of mollusc shell: acid-soluble and acid-insoluble matrix of the limpet Lottia gigantea. Proteome Sci. 2012, 10, 28.
  • 10
    Marie, B., Joubert, C., Tayalé, A., Zanella-Cléon, I. et al., Different secretory repertoires control the biomineralization processes of prism and nacre deposition of the pearl oyster shell. Proc. Natl. Acad. Sci. U.S.A. 2012, 109, 2098620991.
  • 11
    Zhang, G., Fang, X., Guo, X., Li, L. et al., The oyster genome reveals stress adaptation and complexity of shell formation. Nature 2012, 490, 4954.
  • 12
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