J. Mol. Biol. (1991) 220, 687-700

Identification and Characterization of Drosophila melanogaster Paramyosin Javier Vinhs, Albert0 Domingo, Roberto Marco and Margarita Cerverat Departamento de Bioquimica de la UAM and Instituto de Investigaciones BiomMicas de1 CSIC Facultad de Medic&a, Universidad Authoma de Madrid Arzobispo Morcillo, 4, Madrid 28029, Spain (Received

10 October 1989; accepted 5 April

1991)

Paramyosin, a major structural component of thick filaments in invertebrates has been isolated, purified and characterized from whole adult Drosophila melanogaster extracts and a specific polyclonal antibody against it has been prepared. Paramyosin has been identified on the basis of several criteria, including molecular weight, cr-helicity, species distribution, capability of fiber formation in vitro and sequence. We have used the immunopurified polyclonal antibody to isolate eight clones from a lgtl 1 expression library of Drosophila 1 to 22 h embryo cDNA. The largest clone (pJV9) has been sequenced and encodes the coiled-coil region of D. melanogaster paramyosin that is 47% identical to Caenorhahditis rlegans paramyosin. Indirect immunofluorescence in semi-thin sections of adult flies show fluorescence mainly in tubular muscle. Freshly prepared tubular myofibrils decorated with the immunoabsorbed antibody show the A region in the sarcomere as the specific localization of paramyosin. The amount of paramyosin in tubular synchronous muscles of insects appears to be five times higher than in fibrillar insect muscles. There are at least three paramyosin isoforms as shown by isoelectrofocusing separation. The more acidic and less abundant form is phosphorylatrd as shown by 32P in vivo labeling experiments in adult flies. The developmental pattern of expression of Drosophila paramyosin is presented. This mrsoderm-specific protein, immunologically undetectable during gastrulation and early phases of germ band formation, progressively increases during organogenesis to the adult stage. Interestingly, it is also expressed as a major maternal product in the insoluble caytoskeletal fraction of the mature oocyte. Keywords:

Drosophila;

paramyosin;

1. Introduction Bot>h ultrastructurally and biochemically, insect muscle is similar to vertebrate muscle. Historically, insect muscle has been used extensively for investigating the structure and organization of contractile 1984; Armitage et al., tissues (Smith, 1972). Nevertheless, recent molecular and biochemical studies conducted on different types of vertebrate muscle (for example, see Kedes & Stockdale, 1988) have not yet been fully extended to invertebrate systems, in spite of the interest in systems such as Caenorhabditis elegans and Drosophila melanogaster, with their unique genetic and developmental manipulability. A complete two-dimensional map of the 7 Author

addressed.

to whom all correspondence should be

phosphorylation;

isoforms

polypeptides enriched in D. melanogaster thoraces and asynchronous indirect muscle fibrils has been published (Mogami et al., 1982), but the functional role of most of these proteins remains to be established. Only a few primary myofibrillar structural proteins have been satisfactorily characterized in Drosophila, at both protein and gene levels: these are actins (Fyrberg et al., 1980; Tobin et al., 1980), myosin heavy chains (Bernstein et al., 1983; Rozek & Davidson, 1983; Kiehart et al., 1989), myosin light chains (Falkenthal et al., 1984; Parker et al., 1985), and tropomyosin (Basi et al., 1984; Bautch et al., 1982). Recently, the D. melanogaster cr-actinin (Fyrberg et al., 1990) has been cloned and sequenced, and genetic and molecular analyses in Drosophila have begun to advance in the understanding of myofibrillar structure, assembly and function (Fyrberg & Heall, 1990). In addition to

688

J. Vinds et al.

these, a few other proteins have been described in insects (Bullard et al., 1973a,b; Hammond & Gall, 1975; Saide et al., 1989), such as paramyosin or projectin, but their fine structural and functional roles remain to be established. In this paper, we describe the purification and characterization of a 107 x lo3 M, protein identified as Drosophila paramyosin. Paramyosin is a major structural component of thick filaments isolated from many invertebrates muscles (Cohen et al., 1970; Levine et al., 1976; Bullard et al., 1973a,b). This protein was first discovered and isolated as a component of the large filaments in unstriated molluscan muscles (Cohen & Szent-Gyorgyi, 1957). The protein has a rod-shaped structure that appears to be fully a-helical from optical rotatory dispersion measurements (Cohen et al., 1970, 1971). Indirect immunofluorescence in Drosophila shows that the protein is present mainly in tubular synchronous muscle. Although some biochemical properties are not identical to those of paramyosins described in other insect systems (Bullard et al., 1973a,b), its amino acid sequence shows a homology to C. elegana paramyosin.

chromatography of the pooled hydroxylapatite fractions in the same conditions as the first DEAE column allowed us to recover essentially pure paramyosin. Fractions containing the protein were pooled and stored at 4°C. The antibody elicitation method used has been described (Springer et al., 1977). Coomassie-stained bands gels of a 107 X lo3 M, protein from 10% polyacrylamide of insoluble cytoskeletal fractions were cut out to obtain a polyclonal antibody in rabbits, which was used to work out the above described purification protocol. Once the purified protein was obtained. it was electrophoresed and cut from the gel again to obtain a purer antibody in rabbits. After homogenization of the protein polyacrylamide slice with 75% Freund’s adjuvant, the suspension was intramuscularly injected into a rabbit. The first immunization used Freund’s complete adjuvant: the injection was repeated weekly with Freund’s incomplete adjuvant. After 5 injections, rabbit serum was tested for anti-paramyosin reactivity. Anti-paramyosin IgG was purified according to the *Johnstone method (Johnstone & Thorpe, 1982). and subsequently immunoadsorbed on an affinity column prepared with the purified protein as described (Tan-Wilson et al., 1976: Van Eijk & Van h’oor, 1976). Paramyosin was purified from Mytilus retractor muscle by the ethanol precipitation technique as described (Levine et al., 1982). The anti-Mytilus paramyosin antibody was prepared following the same protocol.

2. Materials and Methods Unless otherwise specified, reagents used were of chemical analytical grade. D. melunogueter Oregon R was used as standard strain. Embryos and adult flies were collected as described (Karr & Alberts, 1986). Stage 14 oocytes were obtained individually by hand isolation or in bulk using a modification of the method of Petri et al. (1976). (a) Purification

and antibody preparation

All procedures were at 4°C and buffers contained 61 mg soybean trypsin inhibitor/ml and 1 mg antipain/ ml to prevent proteolysis. Paramyosin was isolated from Drosophila extracts as follows: whole adult flies were homogenized l/10 (w/v) in buffer H (50 mM-Tris.HCI, pH 7.0, 100 mM-Pu’aCl, 300 mM-SUCrOSe, 65% Triton X-100). After filtration and centrifugation (10,000 g for 20 min at 4”C), the pellet was resuspended in the same buffer and centrifuged again. The with buffer W material was washed insoluble (10 mw-Tris. HCI, pH 7.0, 140 mM-NaCl), and the extract’ was recentrifuged. After these washing steps, the protein was solubilized overnight in the salt-containing buffer R (10 miv-Tris. HCl, pH 80 and 0.6 M-KaCl). Upon centrisupernatant was dialyzed fugation , the solubilized exhaustively against 10 m&r-Tris.HCl (pH 8.0) and clarified by centrifugation for 30 min. The final supernatant was fractionated through a DEAE-Sephacel column Uppsala, Sweden) equilibrated with (Pharmacia, 10 mw-Tris (pH 8.0). This step separated paramyosin from actin and myosin. Proteins were eluted with a 0 to 65 M-NaCl gradient in the same buffer; paramyosin eluted at approximately 618 M-NaCl. Fractions of the greatest purity were pooled and dialyzed against 30 mM-/I-mercaptoethanol and 10 mM-sodium phosphate (pH 7.5), and chromatographed on a hydroxylapatite column (Bio-Rad, Richmond, CA, U.S.A.) equilibrated in the same buffer. Proteins were eluted with a 001 to 64 M-sodium phosphate gradient. Paramyosin eluted at 615 M-sodium phosphate. A second DEAE-Sephacel

(b) (lircular

dichroism

From the circular dichroism data of purified paramyosin , the a-helix percentage was calculated using CDPROT, a microcomputer program kindly provided by Dr F. Moran (Menendez-Arias et al., 1988). The best adjustment to the experimental data was obtained using approaches based on proteins of known Y-dimensional structure. Using the method of Bolotina et al. (1980), the root-mean-square error obtained was 1.55, with an a-helix content of SO%, 4% fi-structure, 1276 p-turn and 40/; random structure. Using the approach of Chen et al. (1974) the rootmean-square error was 2.51, with 88% a-helix, 3% p-structure and 9% random structure. (c) S’DSlpolyacrylamide gels, two-dimensional protein gel electrophoresis, immunoblot analysis and densitometric scanning of the gels SDS/lO~/, polyacrylamide gel electrophoresis was performed as described by Laemmli (1970). For 2-dimrnsional gels, embryos, thoraces or whole adult flies were homogenized directly in isoelectrofocusing sample buffer. Thoraces were obtained after acetone-freeze-dry microdissection of adult flies? as described by Mogami et al. (1982). The separation of solubilized proteins by isoelectrofocusing was carried out according the the method of O’Farrell (1975). Separation in the 2nd dimension was by molecular weight on SDS/lO”/o polyacrylamide gels. Electrophoretic transfer onto nitrocellulose sheets or Immobilon polyvinylidene difluoride transfer membranes (Millipore, Bedford, MA. U.S.A.) was performed essentially as described (Towbin et al., 1979). After incubation of the nitrocellulose sheets with rabbit, polyclonal antibody, the reaction was visualized by treatment with peroxidase-labeled goat anti-rabbit antibody (NORDIC. Tilburg, The Netherlands) followed by incubation with substrate. Scanning of polyacrylamide gel electrophoresis samples was performed on a Hoefer GS 300 Transmittance/

Identijication

of D. melanogaster Paramyosin

Reflectance Scanning Densitometer (Hoefer Scientific Instruments. San Francisco, CA, U.S.A.). Tubular myofibrils (purified as described below), 4.5 to 8.25 pg per lane, and fibrillar myofibrilis, 7.0 to 9.5 pg per lane, were loaded in 7% polyacrylamide gels and Coomassie blue stained after electrophoresis. The resultant scanning data were analyzed in the GS 360 Data System PC version (H.S.I.). (d) Preparation

oj tu.bular and jbrillar

Jibers of Drosophila

Fibrillar fibers were isolated from adult wild-type flies as described @aide et a,l., 1989). Tubular fibers were prepared from 3rd-instar larvae using a modification of this method. Approximately 20 g of larvae were suspended in 140 ml of Hodges/EGTA buffer (01 M-KCl. 0001 M-i&$!,, 2.5 mmEGTA. pH.7) containing 65 mM-phenylmethy1sulfonyl fluoride and 61 mg soybean trypsin inhibitor/ml. Larvae were homogenized and filtered through nylon mesh (30 threads/inch) to remove large debris and the filtrate was centrifuged at 800 g for IO min in a Beckman TJ-6R centrifuge. Pellets were resuspended in the same buffer and centrifuged to remove soluble proteins. Following this centrifugation, pellets were resuspended and filtered through a Nytex screen with a pore size of 50 pm diameter. The material remaining in the Nytex was resuspended in a small volume of Hodges/EGTA buffer. Samples were loaded in 38 ml centrifuge tubes onto 2 layers of 18 ml of 1.7 M and 5 ml of 683 M-sucrose in the same buffer. Tubes were centrifuged in a SW28 Beckman rotor at 10,000 revs/min for 30 min to sediment tubular myofibers. This fraction was resuspended in 683 M-sucrose/Hodges/EGTA buffer and reapplied to a 2nd discontinuous gradient prepared in the same tubes with 7 ml layers of 25, 2 and 1.7 M-sucrose Hodges/EGTA buffer centrifuged at in and 22,000 revsjmin for 1 h. Pure tubular fibers were removed with a Pasteur pipette from t,he interface between the 2.5 M and 2 M-XUCrOS~ layers. (e) Immunojluorescence

microscopy

Adult flies were fixed using the microwave fixation method (Login, 1978; Login & Dvorak, 1988). After fixation and inclusion in Tissue-Tek (Miles Scientific, saperville, IL, U.S.A.), flies were frozen and semi-thin sections prepared in a cryotome. Microwave fixation has been reported to be a very effective method for maintaining morphology, with better preservation of the antigenicity of biological samples. Indirect immunofluorescence staining of frozen sections and isolated myofibrils followed standard procedures (Pate1 et al.. 1987) using commercial fluorescein or rhodamine-labeled second antibodies at an appropriate dilution (goat anti-rabbit antibodies. Nordic). (f) Library screening and nucleotide sequencing The D. melanogaster 0 to 22 h embryonic cDNA library in the expression vector lgtl 1 was kindly provided by Dr Francisco ,Jimenez, originally prepared from randomprimed cDNA by Dr B. Hovemann. Approximately 1 x lo5 plaque-forming units from the expression library were plated on Escherichia coli Y1090 in soft LB agar containing 10 mr+r-MgCl, on LB agar (Young & Davis, 1983). After incubation at 42°C for 4 h, plates were transferred to 37°C and overlayed with nitrocellulose filters (Schleicher $ Schuell, Dassell, W. Ctirmany) soaked in 10 mM-isopropyl-fl-D-thiogalacto-

689

pvranoside and blotted drv on filter paper immediately prior to use. Filter orientation was marked using a toot,hpick. Plates were incubated for 3 h at 37°C before removing the filters, and stored at 4’C during filter processing. Filters were saturated in NET buffer 5 mu-Tris.HCl. pH 7. (615 mM-NaCI, 5 mM-EDTA. 605% Triton X- 100, 0.25 y0 gelatin) overnight at 4 *C and immunostained as described for Western blots. Phage plaques giving a positive signal were picked using the wide end of a Pasteur pipette and successively rescreened t,o purify recombinants. Lambda recombinant cDXA inserts were transferred to plasmid Bluescript, and one (pJV9) was sequenced by shotgun strategy (Bankier et al., 1987) in M13mpl9. employing the dideoxy protocol and Sequenase (United States Biochemical, Cleveland, U.S.A.). The Staden Microcomputer Package Software (Staden, 1982) was used to analyze the data, with an average number of 5 gel characters per consensus character. Dotty Plotter computer program for the Macintosh (Don Gilbert. Biocomputing Office, Indiana University. Bloomington. IN) was used to obtain the plot shown in Fig. 2 (below). (g) In vivo labeling of adult flies A 1 g sample of adult flies was collected and labeled with 2 mCi of 32P (orthophosphate in dilute hydrochloric acid from Amersham). The radioactivity was soaked in a filter paper inserted in a bottle with the flies. After 24 h, flies were collected and acetone-freeze-dry microdissection of adult flies thoraces was performed. Samples were prepared according to the method of Mogami et al. (1982).

3. Results (a) Purification

of paramyosin JIY homogenates

from adult

The small size of D. melanogaster adults makes it difficult to obtain large quantities of any single tissue, thus purification of muscle proteins had to be initiated

from

quantitatively,

whole

flies.

Nevertheless,

at

least

muscle is the principal tissue type. In Figure 1, a flow chart summarizes the purification procedure (see Materials and Methods). Until the salt-extraction step (buffer R), paramyosin is recovered in the insoluble fraction. The solubilized protein is further fractionated by DEAE-Sephacel and hydroxylapatite chromatography. As demonstrated by quantitative densitometry, paramyosin obtained by this procedure was more than 900/, pure. Only low molecular weight contaminants were present (Fig. l(b), lane 6), which were separated (although not completely resolved) in a second DEAE-chromatography the run of pooled hydroxylapatite fractions. A new pool of the 107 x lo3 Jf, protein peak (shaded fractions in Fig. l(a)) was more than 95% pure showing only a single protein band by Coomassie blue staining (Fig. l(b), lane 7). Antibodies to the 107 x lo3 M, protein were first obtained by injection of a band of this molecular weight cut from SDS/polyacrylamide gels of the cytoskeletal pellet (i.e. the pellet obtained after homogenate centrifugation). Once the protein was purified, a new polyclonal antibody was obtained.

690

J. Vinds et al.

(al

(b)

(cl

Figure 1. Scheme of paramyosin purification from Drosophila adult flies. In (a) the chart flow is presented. Buffers and fract,ions are explained in Materials and Methods. Shaded areas in chromatographs represent the fractions where paramyosin is detected. In (b) SDS/lo% polyacrylamide gels, stained with Coomassie blue, of representative fractions from the purification are shown. Lanes: 1, homogenate; 2, cytoskeletal pellet (pellet, after centrifugation of the homogenate); 3, fraction solubilized in the salt-containing buffer R; 4, fraction loaded in the DEAE-Sephacel; 5. 6 and 7 show t#he patterns of proteins in the pools of the first DEAE-Sephacel, hydroxylapatite and 2nd DEAICSephacel chromatographs, respectively (pooled fractions from the shaded areas in the scheme). In (c) SDSjlO~~ polyacrylamidr gel stained with Coomassie blue (lane 1) of the cytoskeletal pellet (equivalent to lane 2) and the corresponding Western blot of the same fraction incubated with anti-paramyosin antibody (lane 2). Pi/Na, sodium phosphate gradient,.

The purification procedure was followed using the polyclonal antibodies. In Figure 1(c) (lanes 1 and 2), the crossreaction of the cytoskeletal pellet with the antibody can be seen. Paramyosin is a dimer in native conditions as has been demonstrated previously (Bullard et al., 19733). The analysis in polyacrylamide gels of the 107 x lo3 M, protein from Drosophila cytoskeletal pellet in denaturing conditions in the absence of /?-mercaptoethanol, after Western blots, suggest that paramyosin dimer is stabilized not only by hydrophobic forces but also by disulfide bridges (data not shown). Under the same conditions the purified 107 x lo3 M, protein runs as a dimer, trimer, tetramer etc.! indicating its tendency to polymerize. (b) 1dentiJication of the 107 x IO3 M, protein Drosophila paramyosin

as

Paramyosin (Bullard et al., 19733; Levine et al., 1976, 1982; Elfvin et al., 1976), recently cloned in the nematode C. elegans (Kagawa et al., 1989), is a protein associated with the thick filaments in invertebrate muscles, forming the “core” of these filaments and possibly controlling their length. Paramyosin is a 90 to 100% a-helical, coiled-coil protein composed of two identical polypeptide chains rich in glutamic acid with a molecular weight of 210 to 230 ( x 103) M,. In previous studies (Elfvin et al., 1976; Levine et al., 1976), it has been reported that paramyosin in homogenates from arthropod

and mollusc muscles is composed of two identical polypeptide chains. The Drosophila 107 x lo3 M, protein purified here has been identified as paramyosin by several criteria. (1) Ultraviolet circular dichroism spectra of the purified protein show two negative troughs at 208 and 220 nm and are very similar to the circular dichroism spectra of proteins that exist largely in a-helical conformation. Using two different methods (Bolotina et at., 1980; Chen et al., 1974), we have obtained a very high a-helix content for the purified protein, from 80 to 88%, very close to the percentage described for paramyosins (Bullard et al., 19736). (2) The presence of a similar crossreactive protein was investigated in different species. Cytoskeletal proteins and the corresponding soluble fractions from vertebrate muscle (striated and smooth muscle from cat, chicken and rat) and several invertebrate extracts (Artemia, Drosophila) or isolated muscle homogenates (Astacus, Helix, Locusta, Lumbricus, Mytilus) were analyzed in SDS/polyacrylamide gels. Control gels were run with purified Drosophila 107 x lo3 M, protein and intact and degraded myosins to eliminate the possibility that breakdown products of myosin heavy chains, which might have a mobility similar to that of paramyosin, crossreacted with this antibody. Crossreactivity with bands of similar molecular weight was found in all arthropod, mollusc and annelid species analyzed (Artemia,

Drosophila,

Astacus,

Lumbricue,

Helix.

Zdent@cation

of D. melanogaster

Paramyoein

10 20 30 40 50 60 70 80 CTG~XCAA~TGT~'CA~CAPW~AWW~ LRKLLEDVHLESEETTLLLKKKHNEIITDFOEQVEILTKN 130 140 150 160 170 180 190 200 ACAGAX~~GA KARAEKDKAKFQTEVYELLSQIESYNKEKIVSEKHISKLE 250 260 270 280 290 300 310 320 G~CATCTCCGAGCPCMCCn;MGATCC;AGCACCTPMC VS I SE LNVK I E E LNRTV I D I S S isoRS RL 370 380 390 400 410 420 440 CwmmccGc-cB L KVQLDTVSF S KSQVI SQLE DARRRLE 490 500 510 520 530 540 550 560 CA~TCGAA~GTAA~CGAGGCCCGCA-CAATGCCGA~~~~C E ES5; S E 6;. R I ;70L E RsS; Ii Q Ilo" I E62; D S 6;. R N troL

691 90 100 CGAAAAAC

110

120

210 220 230 240 TCDFOIYX;(;AGMGCAUTCCMACTGGAG 330

340

SQENI 450

350

360

DEDRRR.S 570

II VQD 470 480 GTfXCTCWITGCTCGAAX~TCTG LL E S S L 580 590 600

L V 6;.

N

TCCGTCGCAAGTAC-CGCA=-CATCBTC-VT -CCCKWNSEVAARAEEVEEIRRKYQVRITELEEHIESLIVKVNN 730 740 750 760 770 780 790 800 CT-CAAGTCLEKHKTRLASEVEVLIIDLEKSNNSCRELTKSVNTLEKHN 850 860 870 880 890 900 910 920 -cmACGAGAC-GCGAC-GACCTCGTCCP VELKSRLDETIILYETSQRDLKNKHADLVRTVHELDKVKD 970 980 990 1000 1010 1020 1030 1040 M~~~CPGCICC~~C~~~CTG NNNQLTRENKKLGDDLREAKGAINELNRRLHELELELRRL 1090 1100 1110 1120 1130 1140 1150 1160 GAGAAcGAGcC--Aw2c~Gc~GT ENE RD E LTAAYKEAEAGRKAE EQRGQRLAADF 1210 1220 1230 1240 1250 1260 1270 1280 CGTCn;GCCCA~---X-CC= RLAEKDEEIEAIRKQTSIEIEQLNARVIEAETRLKTEVTR 1330 1340 1350 1360 1370 1380 1390 1400

:60LTK

A

D 700

A

810

820

830 840 azcTcGAGAA~cAAC

930

940

950

960 C

1050

1060

1070

1080

1170

1180

1190

1200

1290

NQYRHAER 1300 1310

1320

A

T S 710

N

Q N 720

ATCM-TCCAGA-TGTCGCTfXA~CAACATCGA~

IKKKLQIQITELEMSLDVANKTNIDLQK

(a 1

0

200

600

400

800 L

J

Base window: 25 Stringency: 10 Points: 372 / I I 1 1 I I 1 I I

I I I1

1 I I I I,

I I r I I 1 I I I,

I I I I I1

-

1 I I , I 1 v-0

C. eleguns paramyosin (0 to 867) (b) Figure 2. (a) DNA sequence and the derived protein sequence of the pJV9 clone from Drosophiln paramyosin. peptide dot,-plot comparison between the sequence of Drosophila and C. eleguns paramyosin is shown.

Mytilus and Locusta) but not in striated, smooth vertebrate muscle or other types of vertebrate tissue (not shown). Furthermore, a paramyosin polyclonal antibody prepared from the mollusc Mytilus

(b) The

anterior byssus retractor muscle (for which a maximum paramyosin/myosin ratio has been determined) crossreacts with Drosophila paramyosin, although at low titers. According to these results, a

J. Vids

similar cross-reactive polypeptide, not found in vertebrates, is present in all invertebrates tested, as could be expected for paramyosin. (3) Finally, cDNA expressing a polypeptide recognized by the polyclonal antiserum against the purified 107 x IO3 M, protein has been isolated by screening a Drosophila embryonic cDNA expression library. From 1 x lo5 independent recombinants, eight antibody-positive plaques were identified. All eight cDNA clones are closely related, as shown by crosshybridization and by immunological analysis of their fusion protein expression products in E. coli. The eight clones, with inserts ranging from @8 to 1.4 kbt, were subcloned in the plasmid pBluescript II. Clone pJV9, the longest of the eight, was sequenced by random DNA strategy (Bankier et al., 1987) using M13mp19. The sequence of this clone (Fig. 2(a)) has 468 codons. One reading frame did not contain stop codons and was in agreement with the known codon preference of Drosophila (Aota et al., 1988). We have used this partial amino acid sequence to search the Swissprot data bank using the FASTA program (Lipman & Pearson, 1985; Pearson, 1990). The protein with the best score in this search is C. elegans paramyosin with 47% identity and a Schistosoma mansoni paramyosin 439 amino acid fragment with 35% identity, followed by a group of 20 myosins with sequence identities ranging from 34% to 19%. As further proof of the homology between C. elegans and D. melanogaster paramyosins, the dot plot in Figure 2(b) is shown. A diagonal beginning at 102 amino acids from the initial methionine of the C. elegans sequence stands out clearly, indicating that this 468 amino acid fragment corresponds to an internal part of D. melanogaster paramyosin . (c) Characterization

of Drosophila

paramyosin

The electrophoretic mobility in SDS/lo% polyacrylamide gels of Mytilus paramyosin is slightly different from that of Drosophila paramyosin. Estimates of Drosophila paramyosin molecular weight were made from homogenates prepared with fly thoraces in Laemmli (1970) buffer using standard curves according to the method of Weber & Osborn (1969). A molecular weight in the range of other insect paramyosins was observed (107 x lo3 M,), as reported earlier by Bullard et al. (1973a,b). As described for Mercenaria paramyosin (Stafford & Yphantis, 1972), Drosophila paramyosin molecular weight decreases slightly during purification, in our hands, after solubilization with salt. An extensive study and classification of Drosophila muscle proteins was published by Mogami et al. ‘(1982), which describes 186 proteins identified by Coomassie blue staining of two-dimensional gel electrophoresis of fly thoraces and indirect flight muscles. In Figure 3, the Coomassie 7 Abbreviation

used: kb, IO3 bases or base-pairs.

et al. PH 7

-

6

6

Mr x 10-3

I80

II6

64

(0)

(b)

Figure 3. Two-dimensional electrophoresis and antiparamyosin immunoblot analysis of Drosophila thoraces. (a) Coomassie blue-stained electropherogram. Arrowhead the indicates 3 paramyosin isoforms position. (b) Immunoblot analysis of the same sample.

blue staining and the corresponding immunoblot of two-dimensional gels of adult tly thoraces are presented. Small spots of lower mobility appear occasionally and seem to be related to degraded fragments of the protein, as different sample preparations show a large variability in these spots. The 19,20 and 21 myofibrillar proteins described by Mogami et al. (1982), with pI values of 6.10, 6.16 and 6.23, crossreacted strongly with the anti-paramyosin polyclonal antibody. These three protein spots, are, therefore, identified as isoforms of paramyosin. The Drosophila protein shows some differences from lower invertebrate paramyosins. Its trypsin degradation pattern differs from that of Mytilus as the latter is more sensitive to paramyosin, trypsin than is Drosophila paramyosin. Digestion of 20 mg of purified Mytilus paramyosin with @04 mg of trypsin for 30 minutes at 37°C yielded four fragments of 48, 465, 4@5 and 39.5 ( x 103) M,, with 008 mg of trypsin, two fragments of 455 and 385 ( x 103) M, were found and @12 mg resulted in complete digestion. The same amount of Drosophila digested by @4 mg and @6 mg of paramyosin trypsin

under

the

same

conditions

gives

three

different protease-resistant fragments of molecular weights 70, 40 and 17 ( x 103) M, (data not shown). Furthermore,

while in our hands purified

Mytilus

paramyosin produced paracrystals in the presence of magnesium in the conditions described for Mytilus (Levine et al., 1982) and for Lethocerus (Bullard et al., 19736), Drosophila paramyosin assembly gave fibers at low pH values regardless of the presence of magnesium, as described below. (d) Drosophila

paramyosin

polymerizes

forming

in vitro $laments

Drosophila paramyosin dialyzed Once solubilized, against water can be visualized on electron micro-

Identijcation

of D. melanogaster Paramyosin

Figure 4. Drosophila paramyosin polymerizes forming in vitro filaments. As described in Results, the purified soluble protein was dialyzed against destilled water and pelleted on a microscope grid covered with carbon film, stained with 1 O/Ouranyl acetate and examined in a Philips electron microscope. Bar represents @5 pm.

scope grids (using 1 o/o uranyl acetate on negative staining) as long, well-formed filaments with a diameter of 10 to 15 nm (Fig. 4). When paramyosin is dialyzed overnight against 50 miw-sodium phosphate buffer (pH 55) these filaments show side-byside aggregation. Filament formation and aggregation is time and pa-dependent. No filaments were visualized by negative staining with 1 o/o phosphotungstic acid when the pH is maintained at 7.5 during dialysis and staining. To test whether or not pH changes are the factor responsible for these differences, the purified protein was dialyzed against low ionic strength buffers at different pH values, using phosphotungstic acid at the same pH as negative staining reagent. Well-formed filaments are visualized only when the pH used is lower than 6, but at pH 65 protofilaments are visible.

(e) Immunolocalization

of Drosophila the adult jZy

paramyosin

in

Indirect immunofluorescence in semi-thin sections of adult flies and isolated myofibrils has been performed for paramyosin localization. Although all insect muscles are striated, in Drosophila, as in other insects, two types of muscles can be distinguished, fibrillar and tubular (Miller,

693

1965; Campos-Ortega & Hartenstein, 1985). Fibrillar muscles are involved indirectly with flight (IFM, Indirect Flight Muscles), contracting very rapidly and asynchronously. They are localized in the thorax and their sarcomeric organization and appearance are similar to those of vertebrate striated muscles. On the other hand, all larval and the majority of adult muscles are tubular, more synchronously fragile mechanically, and contracting, with contraction frequencies similar to vertebrate muscles. In the adult fly, tubular muscles are found in the visceral musculature present around the internal organs within the body cavity and in the hypodermal muscles localized in the inner part of the cuticle all along the body, as well as in certain muscles distributed in the head, legs, thorax and abdomen (Crossley, 1978). Figure 5 shows the fluorescence present in different semi-thin sections of the imago. Controls without the first antibody gave no background fluorescence. In Figure 5(a) and (b), fluorescent and phase-contrast images of hypodermal muscles present in the subcuticle of two different segments of the imago are shown. As seen in the Figure, a fluorescent banding pattern appears when incubated with a paramyosin antibody. Furthermore, the fluorescence appears associated with the ventral and tergosternal muscles (also hypodermal muscles) inserted in the ventral part of the Drosophila abdomen, forming banding patterns transversal to the cuticle (data not shown). In the leg (Bullard et al., 19733; Crossley, 1978; Sandborn et al., 1967) tubular muscles (Fig. 5(c) and (d)) and the intestinal tract (Fig. 5(e) and (f)), the banding pattern is also evident. We conclude that paramyosin polyclonal antibodies in semi-thin sections of D. melanogaeter stain muscle heavily. More specifically, the protein is present mainly in tubular muscle. Only low levels of fluorescence appear associated with fibrillar muscles present in thorax. To understand the display of paramyosin within the sarcomere, freshly prepared myofibrils of larval muscle (no fibrillar muscles exist in larvae) were decorated with the immunoadsorbed antibody. After incubation with paramyosin antibody, fluorescence was associated mainly with the A band in tubular myofibrils (Fig. 6(a) and (b), (c) and (d)). Fluorescence is less intense in the middle of the dark band, suggesting that less protein may be present in the M band region, as has been described for paramyosin in other insects (Elfvin et al., 1976). Controls with adult preparations of fibers in which both fiber types, tubular and fibrillar, are present were negative. No fluorescence appears associated with indirect muscle fibers when dilutions of the antibody that stains tubular fibers brightly are used (Fig. 6(e) and (f)). When lower titers are used, indirect fibrillar fibers prepared from adult flies showed associated fluorescence, indicating that this protein is also present in this type of fiber, but at lower amounts than in tubular muscles. This conclusion is supported by additional data presented below.

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Figure 5. Paramyosin localization in semi-thin section of Drosophila. Anti-paramyosin indirect immunofluorescence ((a), (c) and (e)) and phase-contrast micrographs ((b), (d) and (f)) of imago semi-thin sections are presented: (a) and (b) hypodermal muscles present in the subcuticle between 2 different segments of the imago abdomen; (c) and (d) a leg section at tibia1 level; and (e) and (f) transversal midgut section (visceral musculature). Bar represents 75 pm.

(f) Drosophila

paramyosin is present mainly tubular muscles

in

The presence of paramyosin in Drosophila homogenates is readily detected with our antibody by immunoblotting. In the cytoskeletal pellets, about 95% of the total protein is paramyosin. To make a direct estimation of the quantity of paramyosin in the tubular myofibrils with respect to fibrillar muscles, we have developed a procedure for isolating pure tubular myofibrils from Drosophila larvae (Fig. 7(c)), while indirect flight muscle fibers from adult flies were obtained by published methods

(Saide et al., 1989). Tubular fibers are obtained from larvae to avoid contamination with fibrillar muscle, absent at this stage of development (Crossley, 1978). Paramyosin can be directly quantified by densitometry of the 107 x lo3 M, band in SDS/ polyacrylamide gels, as no other proteins of the same molecular weight are present in two-dimensional gels of these purified myofibril samples (data not shown). Much higher amounts of paramyosin appear in tubular samples than in fibrillar ones (Fig. 7(b) and (d)), as could be expected from the immunofluorescence results summarized above. The

Identification

of D. melanogaster Paramyosin

695

Figure 6. Paramyosin is associated with the A band in Drosophila tubular muscles. The Figure presents antiparamyosin indirect immunofluorescence ((a), (c) and (e)) and the corresponding phase-contrast micrographs ((b), (d) and (f)). Two different tubular

myofibrils

((b) and (d)) of Drosophila larvae show fluorescence associated with A bands (ser

scale in (a)). Adult fibrillar myofibrils ((e) and (f)) do not stain at the antiserum dilution used in this experiment but the tubular

fiber does. Bar represents 25 pm.

amount’ of paramyosin is five times higher in tubular than in fibrillar myofibers (Fig. 7(b) and (d), arrows). Levine et al. (1976), by scanning of purified myosin and paramyosin run in SDS/polyacrylamide gels, determined the range of protein loadings over which Beer’s law holds, as well as the differences in dye binding of both proteins. Using their correction factor we made an estimation of the molar ratio of both proteins (myosin to paramyosin) in Drosophila tubular and fibrillar muscles. The obtained values were 6 to 1 in tubular muscles and 34 to 1 in the fibrillar ones. In agreement with these results, immunoblotting of dissected thorax, abdomen and head shows that paramyosin is present in relatively higher amounts in abdomen and head, in which the majority of the tubular muscles are found, than in thorax (data not shown). The analysis of two-dimensional gels of both fiber types were performed using ampholytes that extend the isoelectrofocusing zone from p1 5 to pI 6.5, to improve the resolution of the paramyosin isoforms. The comparison of both muscle preparations and

corresponding immunoblots indicates t’hat more than one isoform, in addition to their different relative amounts, is present in each type of muscle (data not shown). Further study is needed to clarify the number and relative amounts of isoforms in Drosophila fibrillar and tubular muscles. (g) The more acidic paramyosin phosphorylated

isoform

is

Several groups have shown that paramyosin can be phosphorylated in vitro (Krause & Munson, 1982; Schriefer & Waterston, 1989), but no information exists on in vivo phosphorylation patterns. 32P-labeling experiments with adult Drosophila flies were performed in vivo (see Materials and Methods). Several proteins were heavily labeled, notably the well-known myosin light chain (Toffenetti et al.. 1987; and Fig. 8(b), bottom right corner). The more acidic form of paramyosin, protein 21 according to Mogami et al. (1982), was phosphorylated as deduced from two-dimensional gel analysis of thoraces (Fig. 8, arrows). The relative amount of’ this form, as shown in the Figure. is much lower

J. Vinds et al.

Figure 7. Relative amounts of paramyosin in tubular weTsusfibrillar muscles. In (a) and (c) phase-contrast micrographs of purified fibrillar (a) and tubular myofibers (c) used in the experiment are presented. Bars represent 100 pm. In (b) and (d) the corresponding SDS/polyacrylamide gel lanes and their densitometric scans are shown. Arrows indicate paramyosin peaks.

than

the

other

conclude from less-abundant phosphorylated

two

more

these data that

basic

isoforms.

We

the more acidic

and

form of Drosophila in viva.

(h) Paramyosin

paramyosin

is

expression during development of Drosophila

Analysis

electrophoresis of Figure 8. Two-dimensional ‘*P-labeled adult fly thoraces. (a) The Coomaasie bluestained gel; and (b) autoradiography of the dried gel. Broken lines in (a) indicate the situation of selected phosphorylated spots in (b); e.g. myosin light chain at the bottom right corner. Arrows point to the more acidic paramyosin isoform.

of insoluble

cytoskeletal

proteins

from

various stages of Drosophila development by immunoblotting shows that paramyosin is present at two distinct periods during development (Fig. 9). (1) A protein of the same molecular weight crossreacting with the anti-paramyosin antibody is present in relatively high amounts in the cytoskeletal pellet of mature oocytes, disappearing progressively during early development and becoming virtually undetectable after three hours. The presence of this protein in relatively high amounts has been verified using preparations of bulk-isolated and individually hand-isolated stage 14 oocytes. The role and forms of this musclespecific protein in oocytes and early embryos are under study. The protein is not present or only present at low levels during gastrulation and germ band extension stages. i.e. from three to eight hours postfertilization. (2) Paramyosin increases in amount progressively during middle and late embryogenesis, starting from around ten hours postfertilization. The protein reaches its maximum level at the adult stage. Whole-embryo staining indicates that it is entirely

Identijkation

of D. melanogaster Paramyosin

(a)

116-a4-58-r 485-

36.5,

101

3

5

7

9

11

13

15

17

Ll

L2

Figure 9. Paramyosin is present at 2 different stages of Drosophila development. Coomassie Blue staining of the

SDS/lo% polyacrylamide gel (a) and the corresponding anti-paramyosin immunoblot (b) of cytoskeletal insoluble fractions from various stages of Drosophila development. In (a) and (b), lane numbers correspond to hours ( f 1) of embryonic development. 00, oocytes; Ll and L2, 1st and 2nd instar larvae. respectively.

confined to mesoderm. Immunoperoxidase staining of whole embryos from late stages of development detects high levels of paramyosin in all muscle groups, pharyngeal, somatic and visceral musculatures (data not shown). The antibody does not stain the epidermis or the central nervous system.

4. Discussion Purification and characterization of native paramyosin from D. melanogaster are significant steps in the systematic study of the molecular properties and the functional role of structural muscular proteins. Drosophila paramyosin purification has been carried out from whole organisms, due to the difficulty in starting with a single tissue in this system. This explains the need for a more complex protocol (Fig. 1) than the standard paramyosin purification from Mytilus retractor muscle by the ethanol precipitation technique (Levine et al., 1982). In fact, ethanol precipitation successfully purifies Mytilus paramyosin, as we have verified, but fails to do so in Drosophila (data not shown). The data presented demonstrate that we have identified Drosophila paramyosin. The polyclonal antiserum immunoabsorbed against the 107 x lo3 M, protein was used to identify and isolate cDNA clones from a Agtll expression library. One, pJV9, codes for 468 amino acids; this sequence is 47% identical to the internal part of C. elegant paramyosin. This homology can be also inferred from the diagonal dot plot analysis in

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Figure 2(b). The protein sequence encoded by the pJV9 cDNA clone starts at amino acid 102 of C. elegans paramyosin. Amino acid sequence analysis shows, as expected, the typical 28 amino acid repeat units described in thick filament coiled-coil proteins, indicating that Drosophila paramyosin is a member of this group. Drosophila paramyosin is less sensitive to trypsin digestion and has a different degradation pattern from Mytilus paramyosin, and is characterized by its in vitro tendency to self-assemble at low pH to form extremely long, unbranched filaments, which, with time, become laterally associated, probably indicating coexist in that these molecules filamentous structures within the muscle cell. The immunolocalization of paramyosin in semithin sections olf adult flies indicates that the muscle protein is present in higher amounts in tubular than in fibrillar muscle (Fig. 5). The fluorescence is always associated with the A band, corresponding to thick filaments, as expected for paramyosin (Bullard et al., 19733; Levine et al., 1982, 1976; Elfvin et aZ., 1976). Differences in the myosin/paramyosin molar ratio between fibrillar and tubular muscles in Drosophila are due basically to their different content of paramyosin. Paramyosin varies from 1 y. of the total protein in fibrillar muscle fibers to 5%) in tubular muscle fibers (Fig. 7). Recently, and in accordance with our results, Chun t Falkenthal (1988), using one-dimensional gels of dissected fibers, detected a protein band of around 100 x lo3 M, in tubular muscle that is not detectable in indirect flight muscles. Although they do not identify the protein, it can only be paramyosin. The myosin/paramyosin value of Drosophila fibrillar muscle is 34, while for tubular fibers it is 6. Levine et al. (1976) classified invertebrate muscles as I, II and III with respect to the length of the thick filament and maximum active tension realized. By comparing the filament length and myosin heavy chain/paramyosin ratios among muscles of different species, they suggest the involvement of paramyosin in length determination and active tension generation. Using their criteria, Drosophila fibrillar and tubular muscles should be included in class I. Fibrillar muscles would have one of the highest values and tubular muscles one of the lowest values in the group but in agreement with the differences in paramyosin content, thick filaments are shorter in fibrillar than in tubular muscles. The finding that the correlation between paramyosin content, Mament dimensions and maximum tension development also holds for the different types of fibers in Drosophila could be used to study the molecular and genetic mechanisms underlying this correlation. The results suggest that differences in the functional role of paramyosin in each type of insect muscle, probably in response to their characteristic specialized requirements, are likely to exist. Visceral and somatic tubular muscle fibers serve in the movement of organs that occupy the body cavities. Their myofilaments vary markedly in their degree

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of organization and in their distribution from the configuration observed in vertebrate skeletal and insect flight muscle (Anderson & Ellis, 1967). Different specific proteins are present in each type of muscle and are probably related to their characteristic structure and function. For example, myosin heavy chain and alkali light chain isoforms specific for tubular and fibrillar muscle are produced by alternative RNA splicing in Drosophila (Bernstein et al., 1986; Falkenthal et al., 1985). Several groups have shown that paramyosin can be phosphorylated in vitro (Krause & Munson, 1982; Schriefer & Waterston, 1989). Some of them have implicated phosphorylation in the regulation of the catch state of molluscan muscle (Achazi, 1979; Cooley et al., 1979; Cohen, 1982). Others, as in the case of C. elegans paramyosin (Schriefer & Waterston, 1989), have implicated phosphorylation in the assembly of thick filaments. To our knowledge, the experimental evidence on which these suggestions are based is lacking. Our results show that the more acidic form, which contributes to the total amount of paramyosin only in a minor way, is phosphorylated in vivo. Furthermore, preliminary results indicate a different distribution of paramyosin isoforms in fibrillar and tubular muscles, suggesting the possibility of isoform specificity, as already demonstrated for myosin heavy and alkali light chains in Drosophila. This different distribution of paramyosin, actin and myosin isoforms probably has functional implications as previously suggested (Wade & Kedes, 1989; Emerson & Bernstein, 1987; Kiehart et al., 1989). It has been shown for C. elegans that paramyosin is the product of only one gene. If the same holds for Drosophila, as preliminary data obtained in our laboratory indicates, isoforms and phosphorylation could all be post-translational modifications. Our preliminary data with Drosophila paramyosins indicate that phosphorylation occurs in both types of muscle tissue. The spatial and temporal patterns of expression of paramyosin have also been studied. Paramyosin is regulated during development, and is detected at two different stages. It is present in oocytes, decreasing in quantity in early embryos (3 h of development, at the beginning of gastrulation). Using crossreactivity criteria, the protein is not again detectable until around ten hours of development, progressively increasing its amount from late embryos to the adult stage. In this second period, the protein seems to be confined exclusively to mesoderm. Immunoperoxidase staining of whole embryos from late stages of development with immunoadsorbed anti-paramyosin showed its presence in high amounts to be restricted to musculature. Experiments are in progress to try and determine whether oocyte isoforms are different from the embryonic, larval or adult ones, as well as their temporal pattern of appearance. Independent of this point, maternal expression in high amounts of a protein specific for a defined type of muscle is of obvious interest in connection with the poorly

known organization of the mature Drosophila oocyte cytoskeleton. A concerted molecular and genetic analysis in Drosophila will be crucial in the clarification of the functions and developmental and physiological regulation of these muscle-specific proteins. We thank Manuel Calleja and Marta San Roman for excellent technical assistance. We are grateful to Amparo Cano for critical readings and suggestions for the manu-

script, Dr Federico Mor&n in circular dichroism spectra and Antonio Fernhndez for photograph preparation. This work was supported by funds from the CSIC, DGICYT and FIS.

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Edited by D. DeRosier

Identification and characterization of Drosophila melanogaster paramyosin.

Paramyosin, a major structural component of thick filaments in invertebrates has been isolated, purified and characterized from whole adult Drosophila...
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