Research Article

Comprehensive survey of p94/calpain 3 substrates by comparative
proteomics - Possible regulation of protein synthesis by p94

Yasuko Ono1, Chikako Hayashi1,2, Naoko Doi1,3, Fujiko Kitamura1,3, Mayumi Shindo4, Kenichi
Kudo5, Takuichi Tsubata4, Mitsuaki Yanagida6 and Hiroyuki Sorimachi1,31Department of Enzymatic Regulation for Cell Functions (Calpain Project), The Tokyo
Metropolitan Institute of Medical Science (Rinshoken), Tokyo 113-8613, Japan2

Department of Applied Biological Science, Faculty of Science and Technology, Tokyo Universityof Science, Chiba 278-8510, Japan
3

CREST, Japan Science and Technology (JST), Saitama 332-0012, Japan

4

Proteomics & Small Molecules Division, Applied Biosystems Japan Ltd., Tokyo 104-0032, Japan

5

KYA TECH Corporation, Tokyo 193-0821, Japan

6

Institute for Environmental and Gender Specific Medicine, Juntendo University Graduate Schoolof Medicine, Chiba 279-0021, Japan

Keywords: calpain, fodrin, iTRAQ, substrate, proteolysis, proteomics, muscular dystrophy
Correspondence: Hiroyuki Sorimachi, Department of Enzymatic Regulation for Cell Functions,
The Tokyo Metropolitan Institute of Medical Science (Rinshoken), 3-18-22 Honkomagome,
Bunkyo-ku, Tokyo 113-8613, Japan
E-mail: sorimach@rinshoken.or.jp
Tel: +81-3-3823-2181
Fax: +81-3-3823-2359
Abbreviations: 2D-LC-MALDI, two-dimensional liquid chromatography-MALDI; LGMD2A, limbgirdle muscular dystrophy type 2A; p94, skeletal-muscle-specific calpain, calpain 3; iTRAQ,
isobaric tags for relative and absolute quantitation.

1

Abstract
Calpain is a family of Ca2+-dependent cytosolic cysteine proteases found in almost all eukaryotes
and some bacteria, and is involved in a variety of biological phenomena, including brain function.
Several substrates of calpain are aggressively proteolysed under pathological conditions; e.g., inneurodegenerating processes, fodrin is proteolysed by calpain. Because very small amounts ofsubstrate are proteolysed by calpain under normal biological conditions, the molecular identities
of calpain substrates are largely unknown. In this study, an extensive survey of the substrates of
p94/calpain 3 in COS7 cells was executed using iTRAQTM labelling and 2D-LC-MALDI analysis.
p94 was used because: 1) several p94 splicing variants are expressed in brain tissue eventhough p94 itself is a skeletal-muscle-specific calpain, and 2) it exhibits Ca2+-independent activity
in COS cells, which makes it useful for evaluating the effects of p94 protease activity on proteins
without perturbing the cells. Our approach revealed several novel protein substrates for p94,
including the substrates of conventional calpains, components of the protein synthesis system,and enzymes of the glycolytic pathway. The results demonstrate the usefulness and sensitivity ofthis approach for mining calpain substrates. A combination of this method with other analyticalmethods would contribute to elucidation of the biological relevance of the calpain family.

2

1 Introduction
Calpain (Clan CA, family C02, EC 3.4.22.17) comprises a family of cysteine proteases whose activity isdependent on an increase in cytosolic Ca2+ [1-3]. In the human genome, 14 genes encode calpain
homologues. They are categorized into two groups, ubiquitous and tissue-specific calpains, based on theirexpression profiles. Because the two ubiquitous calpains, μ- and m-calpain, were the first calpains to be
identified, the term ‘conventional calpains’, was applied to them. They are heterodimers and consist oflarge catalytic subunits (CAPN1 and CAPN2, or μCL and mCL) and smaller regulatory subunits (30K
subunits). As protease enzymes, the μ- and m-calpains have common substrate and inhibitor specificities
but differ in their Ca2+ dependence and preference for peptide substrates over protein substrates.
Unexpectedly, it was shown that a lack of m-calpain, but not μ-calpain, causes embryonic lethality [4-7].
Therefore, despite an activation requirement of Ca2+ concentrations of several hundred mM, theimplementability of which is yet to be demonstrated in vivo, it has been proposed that m-calpain plays afundamental role in the maintenance of life and/or is critical for early development, and is capable ofcompensating for some μ-calpain functions.
Under normal cellular conditions, calpain activation is a tightly regulated process coupled to thehomeostasis of [Ca2+]i. Calpain is activated when [Ca2+]i concentrations increase. The cleavage of target
proteins by calpain often alters their functions, activities, and/or structures, modulating many cellularfunctions. Therefore, identification of calpain substrates and elucidation of the effects of calpain
proteolysis on their functions is important for elucidation of the molecular mechanisms of biologicalphenomena that involve calpain. As calpain is important for normal cell function, many pathologicalconditions are associated with over-activation of calpain consequent to loss of [Ca2+]i homeostasis. For
example, in brain ischaemia, neuronal cell death, cardiac dysfunction, and muscular dystrophies, excessive
proteolysis of substrates by calpain, which is not the primary cause of these pathological states, further
compromises cellular functions and aggravates symptoms [8-13]. These conditions indicate that calpain isa promising target for strategies aimed at ameliorating and/or retarding the progression of the symptoms ofthese diseases [14-16].
On the other hand, only one human pathological condition has been shown to result from genetic loss of
calpain protease activity. Limb-girdle muscular dystrophy type 2A (LGMD2A) is caused by a loss of theprotease activity of the skeletal-muscle-specific calpain p94/calpain 3, which is caused by a pathogenic
mutation of the p94 gene, CAPN3 [17, 18]. Studies on the properties of the p94 molecule and on specificcellular events involving p94, such as apoptosis and myogenic differentiation, have contributed to current
understanding of the p94 molecule, and several proteins have been identified as substrates of p94 [19-21].
Although the role that the p94 molecule plays in vivo is not fully understood, it is possible that thedystrophic muscle phenotype observed in LGMD2A is the result of accumulated losses in cellularresponses to various physiological perturbations because p94 lacks its protease activity. An alternativeubiquitous promoter of CAPN3 was recently identified and the expression of several p94 variants in variousorgans, including the brain, has been reported [22-24], suggesting that p94 is important for tissues otherthan skeletal muscles.
The primary structure of p94 is very similar to that of the catalytic subunits of the μ- and m-calpains;
however, several unique and enigmatic properties distinguish p94 from the conventional calpains. Most
significantly, p94 exhibits very rapid autoproteolytic activity in all protein expression systems examined todate [25, 26]. Although native p94 protein is abundant in protein preparations made from skeletal muscles,
it disappears as a result of autolysis during purification, suggesting that a skeletal-muscle-specificmechanism for stabilizing p94 prevails in vivo [27]. As a consequence of its autolysis, analysis of p94
protease activity using standard methodologies is difficult. To elucidate the aetiology of LGMD2A, data isrequired on the identity of the substrates of p94 and on when and how they are proteolysed. As ourprevious study [28] demonstrated that p94 has substrate specificities very similar to those of the μ- and mcalpains, studies on p94 substrates would contribute to the understanding of brain-specific diseases thatinvolve the conventional calpains.
One of the advantages of using p94 is that Ca2+ stimulation is not necessary for its activation; theconventional calpains have to be activated by an increase of [Ca2+]i concentrations provoked by molecules
such as Ca2+ ionophores, the presence of which affects the entire cell. Using COS7 cells transfected with ap94 expression vector and western blot analyses, we previously detected proteolysis of several proteins,suggesting that p94 proteolytic activity is readily exerted and that the proteins identified are potential invivo p94 substrates [17]. Considering that more than 100 proteins have been identified as in vitro

3substrates of conventional calpains [1], many p94 substrates have not yet been identified because of theconstraints of conventional detection methods.
In this study, quantitative proteomic analysis was performed using the COS7 expression system and the
iTRAQTM method [29-31]. The isobaric tagging of peptides using multiplexed iTRAQTM reagents prior to
MS spectrometry allows identification and quantification of the same peptide derived from different origins,e.g., cancer cells at different malignant states. Proteases represent one of the most direct effectors on theamount of substrate proteins. Therefore, quantitative as well as comprehensive proteomic analysis is
promising for identification of novel protease substrates, although few reports have been published [32].
Several proteins were identified as potential p94 substrates; these included known substrates of
conventional calpains, which supports that p94 and the conventional calpains have common substratespecificities, an indication from our previous study [28]. By western blot analysis, fodrin, one of the few invivo substrates of conventional calpains that is proteolysed during ischaemia, was readily detected to beproteolysed by p94. The net abundance of fodrin determined by its trypsin-digested product did notdecrease in concert with the decrease in unproteolysed fodrin levels, suggesting that the proteolytic productby p94 is relatively stable. These data highlight the usefulness of proteomic analysis as a complement toconventional analytical methods for thorough analysis of proteolytic events elicited by calpain.

4

2 Materials and methods
2.1 Protein expression in COS7 cells and iTRAQTM reagent labelling
Cell culture and transfection (electroporation) methods for COS7 cells, cDNA constructs for expressing
human p94s, wild type (WT) and a protease inactive p94:C129S mutant (CS) were described previously[27]. Seventy-two hours after electroporation, harvested cells (ca. 0.5–1.0 × 107 cells) were lysed in 200 μlof 20 mM triethylammonium bicarbonate (Nacalai Tesque, Japan) by sonication. The cell lysate wascentrifuged at 18,000 × g for 20 min at 4 °C, the supernatant was removed, and protein concentration wasquantified using the DC protein assay (Bio-Rad, Japan).
Seventy-two μg of protein from cells expressing either WT or CS were prepared and diluted to a
volume of 20 μl using 0.5 M triethylammonium bicarbonate containing 0.1% (w/v) SDS. Protein was
reduced, alkylated, and trypsin-digested prior to labelling with iTRAQTM reagent according to the
manufacturer’s protocol (ABI, USA). The digested protein prepared from each cell was divided into two
portions for labelling with iTRAQTM 114 and 116 (WT) or 115 and 117 (CS) (Fig. 1A).
2.2 Peptide fractionation for 2D-LC-MALDI-MS/MS analysis
Two-dimensional peptide fractionation was performed on a DiNa Direct Nano-flow LC system equippedwith a MALDI-plate spotter (DiNa MaP; KYA Tech, Japan ) using a strong cation exchange (SCX) column(HiQ Sil SCX, 0.5 mm inside diameter (i.d.) × 35 mm), a reverse-phase (RP) trap column (HiQ Sil C18-3,
0.8 mm i.d. × 3 mm), and an RP analytical column (HiQ Sil C18-3, 0.15 mm i.d. × 50 mm). Labelled
peptides were pooled, diluted with 2.5 volumes of MilliQ water, and acidified to pH 3.0 using 1 M
phosphoric acid. Half of the preparation was used per injection. Peptides trapped on the SCX column were
eluted using a gradient programme consisting of eight linear segments: 0%–5% solvent B in solvent A(solvent A: 0.1% formic acid, 2% acetonitrile; solvent B: 500 mM ammonium-formate buffer [pH 3.0], 2%acetonitrile) for 10 min, followed by 5%–10%, 10%–15%, 15%–20%, 20%–30%, 30%–50%, 50%–100%,
and 100% solvent B in solvent A, each for 10 min at a flow rate of 4,000 nl/min. Eluent from each gradientsegment was pooled and injected into the trap column and then the analytical column using a gradient of
0%–50% solvent D in solvent C over 60 min (solvent C: 0.1% TFA, 2% acetonitrile; solvent D: 0.1% TFA,
70% acetonitrile) and 50%–100% solvent D in solvent C for 10 min at a flow rate of 300 nl/min. The RP
column effluent was mixed with the MALDI matrix solution (4 mg/ml α-cyano-4-hydroxycinnamic aciddissolved in 70% acetonitrile containing 0.1% TFA and 80 μg/ml dibasic ammonium-citrate) flowing at a
rate of 1,400 nl/min at the outlet, and spotted directly onto the ABI 4800 MALDI-plates in a 24 × 16 arrayat a frequency of two spots per min. Four MALDI-plates corresponding to eight gradient segments in SCX
chromatography (2,736 spots) were analysed.
2.3 MS spectrometry and database search
Mass spectra were acquired in batch mode using an ABI4800 Proteomics Analyser operated by 4000 Series
Explorer Software (V. 3.5; ABI). Measurement conditions were set according to the values recommended
by the manufacturer with some modification; the maximum number of peaks for MS/MS acquisition per
spot, the S/N filter, and spot-to-spot tolerance were set at 10, 100 or 150, and 200 ppm, respectively. For
each MALDI-plate, the instrument was set to scan mass spectra between 800 and 4,000 m/z. Two or threecycles of non-redundant MS/MS scans were performed.
All MS/MS spectra were combined and a single database search was performed against primate protein
sequences in the SWISSPROT database using GPS Explorer software (V. 3.6; ABI) and MASCOT as asearch algorithm. Search parameters included modifications by iTRAQTM at N-terminal residues andinternal K and Y residues, methylmethanethiosulphation of cysteine, and one missed cleavage; themaximum peptide rank was set at 2, and the minimum ion score was set at 85%.
2.4 Data analysis
Relative quantification of identical peptides labelled with four different iTRAQTM tags was performed on
the MS/MS peak list by comparing the areas of four signature peaks at 114, 115, 116, and 117 m/z. Values
were normalized using the sum of values obtained for all tagged peptide pairs in the sample; data sets were
imported into MS-Excel for further analysis. Relative changes in the abundance of individual peptides
were defined as follows (S11x [x = 4, 5, 6, or 7] and R represent normalized iTRAQTM signal values and the5ratio, respectively): if R ≥ 1 (0 < R < 1) for any combination of S115 or S117/S114 or S116, the minimum (theinverse of the maximum) was adopted, representing a fold increase (decrease); otherwise, relative change
was expressed as 1.0, meaning that detected changes were not consistent within internally replicatedcomparisons. Peptides with relative change values greater than 1.10 were subjected to manual inspectionof the quality of their MS/MS spectra. MS/MS spectra without a complete set of four measurable
iTRAQTM tags were excluded from the analysis.
2.5 Western blotting and antibodies
Cell lysates prepared as described in section 2.1 were separated by SDS-PAGE and transferred to PVDF
membranes (Millipore, Japan) as previously described. The membranes were probed with a series ofprimary antibodies in combination with appropriate secondary antibodies conjugated to horseradish
peroxidase, and were visualized using POD immunostaining kit (Wako, Japan). The primary antibodies
were: p94-specific anti-IS2 antibody, which was raised in goats [26-28], goat anti-eEF2 antibody (sc-13004,
Santa-Cruz Biotech), mouse anti-beta-actin antibody (A1978, Sigma, Missouri, USA), mouse anti-vimentin
antibody (V6630, Sigma), goat anti-filamin A antibody (F2762, Sigma), mouse monoclonal anti-fodrin
antibody (clone AA6, Biohit, Finland), and mouse anti-annexin A1 and A2 antibodies (clones 29 and 5,
respectively, BD Biosciences, USA). A polyclonal anti-GMMPR antiserum, which specifically detects the150K fodrin fragment proteolysed by calpain, was elicited using rabbits as previously described [33].
2.5 In vitro translation
In vitro translation experiments were performed as previously described using a rabbit reticulocyte lysate invitro translation kit (DuPont-NEN, UK) and L-[35S]-Met in the presence or absence of 380 μg/ml leupeptin
and E-64 as previously described [26]. Briefly, the translation reaction mixture was mixed directly with anequal volume of SDS-PAGE sample buffer at specified times, and subjected to SDS-PAGE. Gels weredried, exposed to Fujix (Japan) imaging plates and X-ray films, and measured and analysed using a Fujix
Model BAS2000 Bioimage Analyser.

63 Results and discussion
In COS7 cells, overexpressed p94 disappears very rapidly because of its autolytic activity (Fig. 5A, p94)
[26] and simultaneously causes specific proteolysis of other proteins such as calpastatin and fodrin [17].
Calpastatin is a specific endogenous inhibitor of conventional calpains and fodrin is an in vivo substrate ofconventional calpains. To identify proteins targeted by p94 protease activity, quantitative proteomic
analysis of COS7 cells expressing either wild type p94 (WT) or a protease-inactive mutant p94:C129S(CS) was performed using the iTRAQTM technique and 2D-LC-MALDI-MS/MS analysis. Equal amountsof trypsin-digested peptides from each sample were individually labelled using one of four iTRAQTM tags,
which are specifically incorporated into free amine groups. As these tags are isobaric, have the same mass,
and are chemically similar to one another, identical peptides originating from different types of cells were
expected to be present in the same fractions throughout the LC separation procedure [29, 30].
Fragmentation of these peptides by MS/MS analysis simultaneously yields four specific peaks derived fromeach iTRAQTM tag at 114, 115, 116, and 117 m/z; the intensity of the signal corresponds to the amount of
peptides labelled with each tag. Thus, the relative abundance of identical peptides in different populationsof cells, either WT or CS, was determined. Consequently, the amount of the parent protein from which the
peptides were derived can be quantified. Integration of this information facilitates proteomic evaluation of
the effect of p94 protease activity on intracellular proteins of culture cells in which conditions close to, if
not equal to, that of the in vivo situation prevail. Proteins whose cellular abundance has been altered by
p94 protease activity are expected to yield R values greater (down-regulated) or smaller (up-regulated) than
1 (no change), where R represents the ratio of iTRAQTM signals for CS/WT. The comparison between WT
and CS samples was performed using four R values (Fig. 1A).
Using a total of 144 μg of protein extracted from COS7 cells (36 μg labelled with each iTRAQTM
reagent), 2,926 peptides—1,610 of which were unique—were identified from 5,197 MS/MS spectra.
Based on MS spectra of elution profiles of 2D-LC, visual inspection of several MS/MS spectra, andstatistical assessment of iTRAQTM signals (Fig. 2A and B), it was concluded that the experimental
procedure yielded a reasonable amount of MS/MS data with reliability and further analysis of the proteomeof COS7 cells expressing p94 was performed. The identified peptides corresponded to 593 proteins, morethan half of which was represented by a single unique peptide (Fig. 1B). As expected, abundant proteins
were associated with many different peptides: vimentin, 23 peptides; HSP90β, 20 peptides; and β-actin, 14
peptides. Five peptides, four of which were unique, were identified for p94, which illustrates the difference
between the sensitivity of this approach and that of other conventional methods such as western blot
analysis (see subsequent results).
Distributions of modified signal ratios of iTRAQTM 116 (WT-2) to 114 (WT-1) and those of 115 (CS1) to 114 (WT-1) were compared to each other. Replicated WT samples showed that signal measurement
was reproducible (mean = –0.0090, SD = 0.2359) (Fig. 2D). Although most peptides were expressed atsimilar levels in WT and CS samples, the standard deviation was larger for this pair, indicating that
peptides whose abundance differed between WT and CS were present (Fig. 2C). These properties are also
evident in the scatter plots of iTRAQTM signals for each of WT-WT and WT-CS pairs (Fig. 3A and B).
The reproducibility of data for WT samples was in good accordance with the expected accuracy of thismethod [34], while replicated analysis of CS samples resulted in slightly poorer reproducibility than that of
WT (Fig. 3C and D). To minimize false positives, only peptides for which the iTRAQTM signal ratios (Rs)of all possible combinations of WT and CS samples, i.e., 115/114, 117/114, 115/116, and 117/116, changed
in identical directions were considered for further analysis.
Extracted peptides were sorted according to the relative change in abundance (= Rmin if R ≥ 1; = 1/Rmax,
if 1 > R > 0, where R = signal ratio of 115 or 117 to 114 or 116) (Table 1). In several studies conductedusing the iTRAQTM methodology, differences greater than 1.2-fold have been suggested to be consideredsignificant [35]. In the present analysis, where a difference is represented by the minimal fold change, 1.10
was used as an arbitrary cut-off point. MS/MS spectra of these peptides were inspected visually. Proteinsrepresented by peptides with reliable MS/MS spectra and whose signal ratios were decreased by WT p94
expression are listed in Table 2. Representative images of MS/MS spectra for amino acid sequencing and
iTRAQTM signal quantification are shown in Fig. 4.
Among the proteins listed in Table 2, p94 was ranked as the protein with the greatest decrease (averaged
Rmin for peptides = 3.024, n = 5) (Fig. 4D). This is the first quantitative demonstration that the best
substrate for p94 is p94 itself. The data also illustrate that the meaning of a quantitative change of a protein
determined by the abundance of peptide derivatives is different from that detected by other methods such as
western blot analysis. An autolytic decrease in the full-length p94 in COS7 cells was detected by western7

blot analysis, which showed that the amount of WT (addition of 94 kDa, 55 kDa and other smaller bands) is
less than 1/100 of that of CS (Fig. 5A “p94”). On the other hand, the quantity of WT p94 was ca. 1/3 ofthat of CS at the peptide level when measured by the iTRAQTM method (Table 2, Fig. 5D). All five (fourunique) peptides (Fig. 5C “p94”) detected for p94 decreased in WT samples to a similar extent. This large
difference of detected relative WT quantity suggests that most of identified peptides for WT p94 by the
iTRAQTM method are derived from extensively autolyzed fragments of p94 that could not be detected bywestern blot analysis due to small molecular mass and heterogeneity of the autolyzed fragments (Fig. 5D).
Similar decreases of all five peptides detected for WT p94 indicate that upon autolysis, different autolyzed
fragments of WT p94 concurrently become more susceptible to subsequent degradation than unautolyzed
p94 protein.
Some of the proteins listed in Table 2 are previously identified substrates for other calpains [36-39] (seethe “Previous report” column in Table 2). At present, it is not biochemically confirmed that the identified
proteins are direct substrates for p94. Together with the previous observation, however, that p94 and theconventional calpains have some common substrate specificities [28], the present results are indicative formolecular context where p94 directly exerts its activity. The identification of filamin-A and vimentin as
substrates is consistent with the observation that ectopic expression of p94 in several different culture cells
causes morphological defects accompanied by proteolysis of several cytoskeletal proteins, including
filamin-A and fodrin [40]. Western blot analysis detected a slight decrease in filamin-A and vimentin in
WT samples, while the decrease in annexin A1 or A2 was less obvious (Fig. 5A). The proteins listed inTable 2 also include components of translation machinery (eEF1α-1 and -2, eEF-2 and initiation factor 3).
eEF-1α is proteolysed by drosophila calpain [36], and eEF-2, which was shown to be related to p94, wasdecreased in WT samples according to western blot analysis (Fig. 5A). These results strongly suggest that
there is interaction between p94 and the translation machinery, which was further confirmed by thefollowing experiment. When p94 was translated in vitro using the rabbit reticulocyte lysate system, p94
exhibited co-translational autolytic activity during the initial phase; the signal intensity for the full-lengthtranslated product was ca. 1/10 lower than that of the μ-calpain large subunit (μCL) translated in the same
system (Fig. 6A, 0–20 min). The intensity of the 94 kDa band as well as that of other smaller bands
reached a local maximum at about 20 min from the start of translation and then decayed (Fig. 6B). In
contrast, μCL exhibited an almost linear increase in 80 kDa band intensity regardless of the presence ofhigh concentrations of the Ser/Cys-protease inhibitors, leupeptin and E64 (Fig. 6, + or –inh.). If p94 only
had autolytic activity, the amount of p94 would not have decayed, but would have asymptoticallyapproached a plateau at which protein synthesis would have been in balance with autolytic degradation.
Thus, the observed local maximum with subsequent decay indicates that synthesis of p94 protein gradually
decreased, which was probably due to proteolysis of the translation machinery by p94 as shown in
iTRAQTM experiments. These data strongly suggest that p94 is involved in regulation of the eukaryote
protein synthesis system, as is the case for the conventional calpains [36].
Other proteins identified as possible p94 substrates revealed novel aspects of the function of p94.
Fructose-bisphosphate aldolase A, glyceraldehyde-3-phosphate (GAP) dehydrogenase, and triosephosphate
isomerase catalyse successive reactions (fructose-bisphosphate → dihydroxy acetonephosphate + GAP →2GAP → 1,3-bisphosphoglyceric acid) in the glycolytic system, which is highly developed in skeletal
muscle. Another proteomic study using 2-DE and transgenic mice overexpressing p94 identified otherglycolytic enzymes as p94 substrates in skeletal muscle [41], suggesting a role for p94 in the glycolytic
cycle [42]. Importin-7 and the annexins are involved in membrane trafficking. As yeast calpain utilizes amembrane-associated scaffold, a similar mechanism might apply to p94 [43, 44]. Proteolysis ofproteasome regulatory subunit RPS6A by p94 may indicate that the ubiquitin proteasome system is subject
to regulation by calpain, at least in certain cases [45]. At present, the relationship between p94 andthioredoxin is unknown. It has yet to be clarified that cleavages of proteins identified in this study arecharacterized under physiological context as well as biochemically. It is particularly important todistinguish direct p94 substrates from those proteolyzed by other proteases activated by overexpression of
p94. The latter case could non-physiologically happen in COS7 cells, since regulatory mechanisms for p94
activity, which are probably effective in skeletal muscle tissues, may absent here. It should be also noticed
that proteins identified from COS7 cells could be alternatives for their counterparts expressed in skeletal
muscle tissues. Integrated understanding of these issues would reveal physiological behaviour of p94 andits relevance to other cellular machineries with more clarity.
A notable difference between p94 and the other proteins listed in Table 2 is that proteins other than p94,
if represented by multiple peptides, were only decreased by a portion of their peptides. This is probably
because the stabilities of products generated from substrate proteins by p94 differ. Fig. 5C illustrates the8distributions of the identified peptides and the reported proteolytic sites of p94, vimentin, and fodrin.
While autolysis causes down-regulation of p94 as a whole molecule, proteolysis of vimentin would confer
other properties on certain parts of the molecule. This fits a scheme in which calpains do not degrade target
proteins, but modify the functions of targets. Consistent with this hypothesis, proteolysis of α-fodrin was
detected in WT-expressing cells by an antibody against a calpain-specific proteolytic product (Fig. 5B,
anti-GMMPR) [17], but is not by iTRAQTM analysis. Six independent peptides from α-fodrin before andafter the calpain cleavage site were detected by MS/MS (Fig. 5C), essentially none of which showed achange in relative abundance between WT and CS samples. This indicates that fodrin proteolysis by p94
did not change the total amount of fodrin. This quantitative information, which is hard to obtain solely bywestern blot analysis (Fig. 5B, anti-fodrin), suggests that limited proteolysis of fodrin by p94 does notcause immediate degradation of fodrin fragments. Although the possibility of interference at atranscriptional/translational level as a consequence of p94 protease activity cannot be excluded, a more
extreme change in the abundance of target proteins as a whole would have been observed if that were thecase.
Western blot analysis showed that the decrease in identified proteins was rather small compared withthe autolytic decrease in p94 (Fig. 5A). This was probably caused in part by much less efficient protein
proteolysis by p94 than p94 autolysis, which could be because of that COS7 cells but not skeletal muscle
cells were used. Although the expected advantage of our COS7 system is that ectopically expressed
p94WT is the only active calpain present when cells were not otherwise stimulated, the lack of stimuli tospike signalling cascades such as [Ca2+]i regulation or cytoskeletal motility might hamper contact between
p94 and substrates and/or other molecules and enhance autolytic rather than proteolytic reaction.
Levels of several proteins were increased in WT samples (Table 1). This might indicate that proteolysis
by p94 removes factors restricting the stability of proteins. If these proteins are p94 substrates, structuralchanges induced by proteolysis may make them more stable. On the other hand, enzymes that promotedegradation of these proteins may be substrates for p94. Peptides with a fold-change greater than 1.1 were
subjected to manual inspection of their MS/MS spectra. Several peptides had reliable MS/MS spectra, but
few represented parental proteins with consistency. Thus, it was difficult to extract proteins whose increasein abundance was dependent on p94 protease activity. It is also possible that the protease activity of p94
was not responsible for increasing the abundance of other proteins. Identification of such proteins from
COS7 cells was not pursued further and left for the future investigation using different cell culture system.

9

4 Concluding remarks
Quantitative comparison of proteomic data obtained from COS7 cells expressing p94WT or p94CS was
efficient in surveying the change of protein abundance caused by p94 protease activity. The advantages ofthis method over the 2-DE method are not only simplicity and reproducibility, but also the ability tomanipulate high molecular-weight proteins as evidenced by the results presented in Table 2 for filamin-A
and importin-7. Simultaneous identification and quantification by iTRAQTM analysis enabled us to detect
potential p94 substrates that might have been overlooked because of a lack of information and/or analytical
tools. Another advantage of the methodology described in this study is that it enables evaluation of theconsequences of proteolytic events from the abundance of various proteolytic fragments from the same
protein. As calpains have been implicated in modulation of the function of substrate proteins, theabundance of the parts of substrate proteins responsible for their functions does not necessarily decrease
upon proteolysis. The quantitative nature and the unbiased sensitivity of the method will be useful for
identifying the fates of various proteolytic products, which will complement other biochemical and cellbiological studies of proteolysis [46]. This methodology can be applied to all calpain species in various
differential analyses of pathological states, transgenic mice, and developmental stages.

10Acknowledgements
We are grateful to Mitsutoshi Okuno, Keiko Cho, Shinobu Imajoh-Ohmi, Keiko Kurosawa, and Hiroyuki
Fukuda for technical assistance and to all of the “Calpain PT” members for experimental support andvaluable discussion. This work was supported in part by research grants MEXT.KAKENHI 17028055 and18076007 (to H.S.), JSPS.KAKENHI 18770124 (to Y.O.), 18380085 (to H.S.), Research Grant 17A-10 for
Nervous and Mental Disorders from the Ministry of Health, Labour and Welfare, and the Takeda Science
Foundation research grant (to H.S.).

11

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[3]Sorimachi, H., Ishiura, S., Suzuki, K., Structure and physiological function of calpains.
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Azam, M., Andrabi, S. S., Sahr, K. E., Kamath, L., et al., Disruption of the mouse mucalpain gene reveals an essential role in platelet function. Mol Cell Biol 2001, 21, 221320.

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Zimmerman, U. J., Boring, L., Pak, J. H., Mukerjee, N., et al., The calpain small subunit
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14Figure legends
Figure 1. Proteomic analysis of COS7 cells expressing p94. (A) Overview of experimental design. (B)
Number of peptides/proteins identified in the 2D-LC-MALDI analysis. More than half the identified
proteins were represented by a single peptide. Eleven per cent of the proteins identified had more than six
peptides; these included vimentin, several different heat shock proteins, and cytoplasmic actins, which are
known to be abundant.

Figure 2. Statistical profiles of LC-MALDI analysis. (A, B) Error distribution of masses in Da (A) or
ppm (B) between observed and calculated/theoretical molecular masses of 2,801 identified MS/MS spectraout of 5,197 spectra. Dotted lines indicate means and ranges for one and two standard deviations. (C, D)
Modified ratios of iTRAQTM signals of 115 (CS-1) to 114 (WT-1) (C) or 116 (WT-2) to 114 (WT-1) (D).
Values of ratios were modified as follows: Modified ratio = R – 1 (if R ≥ 1), 1/R – 1 (if 1 > R > 0), whereR = iTRAQTM signal ratio.

Figure 3. Scatter plots of iTRAQTM signals (absolute values) between 114 (WT-1) and 115 (CS-1) (A),114 (WT-1) and 116 (WT-2) (B), 115 (CS-1) and 117 (CS-2) (C), and 114 (WT-1) and 117 (CS-2) (D).
Ranges of twofold and fourfold, one-half, and one-quarter are indicated by dotted lines. R2, correlation
coefficient.

Figure 4. Examples of MS/MS spectra that exhibited significant differences in iTRAQTM signal ratios.
(A) MS/MS spectrum of precursor ion of 3,410.5859, which was identified as elongation factor 2 with ionscore 72 and I.C. 99.9% (calculated mass is 3,410.5439, error = 0.0420 [12 ppm]). Representative peaks
are shown and calculated masses are indicated in parentheses. The predicted amino acid sequence is shownin the box, which corresponds to residue 801–829 of human elongation factor 2 with N-terminal iTRAQTM
and Cys-MMTS modification (indicated by *). The small square around m/z = 100 is the area magnified in

15

(B). (B) Magnification of iTRAQTM signals shown in (A). (C–G) Magnified iTRAQTM signals of MS/MS
spectrum (data not shown) identified as filamin A (C), annexin A1 (D), annexin A2 (E), vimentin (F), andp94 (G). The predicted peptide corresponds to residues 309–330 of human filamin A, 29–52 of humanannexin A1, 179–195 of human annexin A2, 36–49 of human vimentin, and 733–748 of human p94.

Figure 5. Comparison of identified proteins by iTRAQTM analysis. (A) Western blot analysis of selected
proteins using specific antibodies. Equal amounts of protein extract from COS7 cells expressing either WTor CS mutant of p94 were loaded for each blot. Closed and open arrowheads indicate the full-length andthe autolysed fragments, respectively, detected by anti-pIS2 antibody. The epitope region is indicated by avertical bar in Fig. 5C. AnxA1, annexin A1; AnxA2, annexin A2. (B) 150K fragment of α-fodrin
proteolysed at the calpain-specific site detected by anti-GMMPR in a protein sample from COS7 cells
expressing WT but not CS (open arrow head). The monoclonal anti-fodrin antibody detected several
fragments of different sizes in addition to full-length unproteolysed fodrin (closed arrowhead) in both theWT and CS lanes. *, A proteolysed 150K fragment specific to the WT sample was also detected by the
monoclonal anti-fodrin antibody, but the signal overlaps with an unidentified band with an almost identicalmobility, which was also present in the CS sample. (C) Positional relationships between previously
determined proteolytic sites and peptides identified in the present analysis. The closed arrowheads indicate
autolytic sites for p94 and proteolytic sites of conventional calpains for vimentin and α-fodrin. Closed,
open, and grey circles indicate the positions in proteins of peptides whose relative abundance weredecreased, unchanged, or increased, respectively, in COS7 cells expressing WT p94 relative to cells
expressing p94:CS. Cys, His, and Asn indicate catalytic amino acid residues in p94. Vertical bars indicate
the positions of epitopes for anti-pIS2 (for p94) and anti-GMMPR (for the calpain-specific 150K
proteolytic fragment of fodrin). Bidirectional arrows indicate the autolytic fragment of p94 and the calpain
specific proteolytic fragment of fodrin, which correspond to the open arrowheads in Fig. 5A and 5B,
respectively. NS, IS1, and IS2 are p94-specific insertion sequences. IIa and IIb comprise the calpain
protease domain. III and IV are C2-like and 5EF-hand motif Ca2+-binding domains, respectively. 1A, 1B,
and 2, coiled coil domains. Sr1-19, spectrin repeat. (D) Relative quantities of p94 protein/peptide by
iTRAQTM method and western blot analysis (WB). WT p94 amounts detected by both methods are16

represented as relative to those of CS (CS = 100). Difference between CS and WT by iTRAQ ([a])
corresponds to portions in WT degraded completely (to amino acid) or at least smaller than the length oftrypsin peptides. Difference between iTRAQ and WB in WT ([b]) corresponds to portions in WT degradedpartially, larger than the length of trypsin peptides but small enough not to be detected by WB. Region [c]mainly corresponds to the 55 kDa band in WB (Fig. 5A “p94”, open arrowhead).

Figure 6. Time course of in vitro translation of p94 and the μ-calpain large subunit (μCL). In vitrotranscribed RNA for rat p94 and human μCL were translated with rabbit reticulocyte lysate and 35S-Met in
the presence (+inh.) or absence (-inh.) of 380 μg/ml leupeptin and E64, and sampled at the indicated time(min). Samples are separated by SDS-PAGE, and exposed to X-ray film (A) and an imaging plate. The
radioactivities of 94 kDa (open triangle) and 80 kDa (closed triangle) bands were quantified using a Fujix
Bioimage Analyser, and plotted against the time of sampling (B). Note that the scales for the Y-axes differ
by a factor of 10.

17

Tables
Table 1. Relative abundance quantified by iTRAQTM analysis
Relative change
No. of peptides
No. of proteins1
No. of proteins2

Increased by p94 (WT > CS)> 1.25> 1.15> 1.10
22
48
76
20
40
61
5
2
15

1

Decreased by p94 (WT < CS)
1.10 <
1.15 <
1.25 <
80
52
29
59
44
22
10
10
3

Total

Number of proteins represented by identified peptides.
Number was obtained using averaged iTRAQTM ratios for all peptides representing a protein.

2

18

307
246
45

Table 2. Selected proteins with CS/WT abundance ratios greater than 1
Identified proteins were ranked based on their increase in CS samples; a high rank corresponds to a high ratio of CS/WT, which implies a decrease in
WT expression.
Rank

Protein

Entry

Swiss
Prot
Acc.
No.

Mr

No. ofTM
Mean No. of
No. of
iTRAQ
TMofunique iTRAQ
peptideswith Rmin Rmin peptides peptides> 1.10Previous
reportTotal Ion Score

Best Ion Score

C.I.%

C.I.%

p94
[26]

μ-, mcalpain

1

p94 / calpain3

CAN3_HUMAN

P20807

94,254

5

3.02

4

5

193.98100.0089.06

100.002

Eukaryotic elongation factor 1-alpha 2

EF1A2_HUMAN

Q0563950,470

2

1.65

7

13

320.85100.00

64.72

99.99

3

Eukaryotic translation initiation factor 3 subunit 7

IF37_HUMAN

O15371

63,973

1

1.48

1

1

60.90

99.98

63.14

99.99

4

26S proteasome regulatory subunit 6A

PRS6A_HUMAN

P1798049,204

1

1.34

1

1

71.36

100.0072.89

100.005Filamin-A

FLNA_HUMAN

P21333

280,608

3

1.31

10

21

824.85100.00144.35100.006

Importin-7

IPO7_HUMAN

O95373

119,517

1

1.28

2

5

197.07100.00100.31100.007

Eukaryotic elongation factor 2

EF2_HUMAN

P1363995,207

3

1.28

11

16

749.70100.00131.15100.008Vimentin

VIME_HUMAN

P0867053,520

5

1.25

23

74

1,694.43

100.00156.66100.00[37]*

9Annexin A1

ANXA1_HUMAN

P0408338,583

2

1.23

2

6

218.54100.00188.19100.00[39]*

10Eukaryotic elongation factor 1-alpha 1

EF1A1_HUMAN

P6810450,141

3

1.22

12

35

910.71100.00194.53100.00[36]

11Annexin A7

ANXA7_HUMAN

P2007352,739

2

1.21

1

5

158.97100.00158.97100.00[39]?

12Annexin A2

ANXA2_HUMAN

P0735538,473

2

1.19

16

55

1,470.31

100.00164.31100.00[39]?

13Fructose-bisphosphate aldolase A

ALDOA_HUMAN

P0407539,289

6

1.18

9

37

726.61100.00123.91100.00[47]?

14

Thioredoxin

THIO_HUMAN

P1059911,606

4

1.16

2

16

127.21100.0066.75

100.0015

Glyceraldehyde-3-phosphate dehydrogenase

G3P_HUMAN

P0440635,922

3

1.15

8

26

779.77100.00214.62100.0016

Triosephosphate isomerase

TPIS_HUMAN

P60174

26,538

4

1.14

6

16

501.27100.00119.92100.00TM

Rmin: the minimum value for the iTRAQ*: proteolytic sites were determined.
?: indicative observations are reported.

signal ratio of 115 or 117 to 114 of 116

19

[36]

[40]

[38]*

Ono, Y. et al. Fig. 1
A
COS cells

p94 expression

Wild type (WT)

p94:C129S (CS)++-p94 autolysis
Substrate proteolysis

Peptide preparation
iTRAQ labeling

114

116

115

117

WT-1

WT-2

CS-1

CS-2

2D-LC/MS/MS

5,197 spectra

Database search

2,926 peptides / 593 proteins

Quantification

R = CS-1, 2 / WT-1, 2 > 1.0 for peptides decreased by p94:WT

B(>6) 11 %(5) 5 %
(4) 7 %
(1) 54 %(3) 8 %

(2) 15 %Ono, Y. et al. Fig. 2

A

B
Average = 0.0289
SD (σ) = 0.0445

350

Average = 16.3
SD (σ) = 24.9

250

300
200

frequency

250

+2σ

-2σ

200

-σ

+σ

frequency

-σ

150

150

+σ

-2σ

+2σ

100

100
50
50

0
-0.2

-0.1

0

error (Da)

+0.1

0
-80

+0.2

C

1000

900

900
Average = -0.0001
SD (σ) = 0.3888

800

+80

Average = -0.0090
SD (σ) = 0.2859

700
-σ

600

+σ

-2σ

frequency

frequency

+40

800

700

+2σ

400

500

200

200

100

100

0

0.5

1.0

Modified iTRAQ ratio (115/114)

+σ

-2σ

+2σ

400
300

-0.5

-σ

600

300

0
-1.0

0

error (ppm)

D

1000

500

-40

0
-1.0

-0.5

0

0.5

1.0

Modified iTRAQ ratio (116/114)

Ono, Y. et al. Fig. 3

AB

x 1/4 x 1/2

6

10

114/1155x2
x4

WT-1/CS-1

4

10

R2=0.9893
3

WT-1/WT-2

4

10

R2=0.9933
3

10

10

2

102

103

104

105

2

10

106

102

103

114 (p94WT-1)

115/117

6
10

x2
x4

CS-1/CS-2

5
10

117 (p94CS-2)

117 (p94CS-2)

10

4

10

R2=0.9715
3

2

102

106

x 1/4 x 1/2

114/117x2
x4

WT-1/CS-2

4
10

R2=0.9704
3
10

10

10

105

D

x 1/4 x 1/2

6

5

104

114 (p94WT-1)

C
10

x2
x4

10

116 (p94WT-2)

115 (p94CS-1)

114/1165

10

10

x 1/4 x 1/2

6

10

103

104

115 (p94CS-1)

105

106

2
10
102

103

104

114 (p94WT-1)

105

106

Ono, Y. et al. Fig. 4

A

eEF-2
y7 (822.37)

1

2

3

4

5

6

7

8

9 10 11

12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29*
SNTGGQAFPQC
VFDHWQILPGDPFDNSSR
801
829

*

5000

y15 (1768.84)

Intensity

4000

3000

b22 (2589.18)

2000

1000

0

y4 (463.23)

0

1500

1000

C

eEF-2

(115)

500

WT-1
(116)

(114)

(116)

CS-2

CS-1

CS-2

(115)

(117)

(117)
600

(114)

WT-1

WT-1

(114)

300

200

3500

annexin A1(116)

Intensity

300

400

3000

D

Intensity

Intensity

400

WT-1

2500

800

WT-1
WT-1

2000

CS-1

(117)

(115)

500

Mass (m/z)

filamin-A

CS-2

CS-1

y27 (3065.37)

y11 (1204.56)

500

B

b25 (2948.33)

y10 (1091.48)

400

200
200

100

100

0

0
114

115

116

117

118

0
114

115

Mass (m/z)

E

400

117

114

115

F

G

vimentin

CS-2

CS-1

CS-2

(115)

(117)

(115)

(117)

500

(116)

400

117

118

p94

800

CS-1

CS-2

(115)

(117)

WT-1

WT-1

WT-1

116

Mass (m/z)

CS-1

WT-1

118

Mass (m/z)

annexin A2

500

116

(116)

(114)

600

300

Intensity

Intensity

Intensity

(114)
300

200

400

WT-1

WT-1

(114)

(116)

200
200

100

100

0

0
114

115

116

Mass (m/z)

117

118

0
114

115

116

Mass (m/z)

117

118

114

115

116

Mass (m/z)

117

118

Ono, Y. et al. Fig. 5

A

kDa

97

WT CS

66
kDa

220

45

WT CS

kDa

WT CS

WT CS

WT CS

66

kDa

45

66

WT CS

97

45

170
30
30

30

116
Filamin-A

p94

B

WT CS

AnxA1 AnxA2

Vimentin eEF-2

WT CS

kDa

220
170

∗

116
anti-GMMPR anti-fodrin

CCys
p94 NS

Vimentinα-fodrin

129

IIa

His334
Asn358IS1 IIb

821aar
III

IS2

IV

466aarHead 1A

1B

2

Tail

2472aar
Sr 1

Sr 2

Sr 3

Sr 4

Sr 5

Sr 6

Sr 9

Sr 10

Sr 11

Sr 18

Sr 19

1 2
EF-handOno, Y. et al. Fig. 6

A

Time 0(min)

p94-inh.

p94+inh.

5 10 15 20 25 30 40

0

5 10 15 20 25 30 40 0

µCL-inh.

µCL+inh.

5 10 15 20 25 30 40

0

5 10 15 20 25 30 40

(kDa)
94
67
43

30

B

400

4000
µCL+inh.

p94+inh.

µCL-inh.

3000

300

p94-inh.
µCL signal

p94 signal

2000

200

1000

100

0

10

20

30

Time (min)40