Cell and Tissue Research

, Volume 337, Issue 3, pp 463–476 | Cite as

Post-transcriptional silencing of the Drosophila homolog of human ZASP: a molecular and functional analysis

  • Clara Benna
  • Samantha Peron
  • Giorgia Rizzo
  • Georgine Faulkner
  • Aram Megighian
  • Giuliana Perini
  • Giuseppe Tognon
  • Giorgio Valle
  • Carlo Reggiani
  • Rodolfo Costa
  • Mauro A. Zordan
Regular Article

Abstract

In humans, mutations in ZASP (the gene for Z-band alternatively spliced PDZ-motif protein) are associated with dilated cardiomyopathy and left ventricular non-compaction. In particular, mutations in or around the Zasp motif seem to be related to myofibrillar myopathy. Thus, “zaspopathies” include symptoms such as Z-line disgregation, proximal and distal muscle weakness, cardiomyopathies, and peripheral neuropathies. In order to understand the role of ZASP in muscle structure and function, we have performed a molecular characterization of the Drosophila ortholog of human ZASP and a functional analysis following the post-transcriptional silencing of the Drosophila gene. Transcriptional analysis of dzasp has revealed six additional exons, with respect to the known 16, and multiple splice variants. We have produced transgenic lines harboring constructs that, through the use of the UAS/Gal4 binary system, have enabled us to drive dsRNA interference of dzasp in a tissue-specific manner. Knockdown individuals show locomotor defects associated with alterations of muscle structure and ultrastructure, consistent with a role of dzasp specifically in the maintenance of muscular integrity.

Keywords

dsRNA interference Muscle Z-line Myopathies ZASP Drosophila melanogaster (Insecta) 

Introduction

ZASP (Z-band alternatively spliced PDZ-motif protein)/Cypher/Oracle (Faulkner et al. 1999; Zhou et al. 1999; Passier et al. 2000) is a PDZ/LIM protein that belongs to the ALP/Enigma family. The various isoforms of ZASP have either three or no LIM domains, but all have an N-terminal PDZ domain. All members of the ALP/Enigma family of proteins have one PDZ domain, but only ZASP and Enigma have three LIM domains. A “ZASP-like motif” (ZM) is present in ZASP and two other members of the family, ALP and CLP-36. ZASP and other members of the family interact with alpha-actinin-2 via the PDZ domain (Au et al. 2004); however, at least for ALP, the ZM is also required for alpha-actinin-2 interaction (Klaavuniemi et al. 2004). The ZM has been proposed to be needed to stabilize and reinforce the interaction with alpha-actinin-2. Recently, this domain in ZASP has been shown to be independently co-localized with alpha-actinin-2 at the Z-line (Klaavuniemi and Ylanne 2006). ZASP is mainly expressed in striated muscles in mammals, but low levels of expression have been reported for several other tissues, e.g., in mice in brain, placenta, pancreas (Faulkner et al. 1999) and lung (Zhou et al. 1999) and in humans in brain, lung, kidney, uterus, and part of the gut (Faulkner et al. 1999; Arimura et al. 2004).

In mice, a total of six alternatively spliced ZASP isoforms have been described that include three cardiac muscle-specific isoforms and three skeletal muscle-specific isoforms (Huang et al. 2003), these can be classified into long or short isoforms. All of the latter ZASP isoforms have both a PDZ and a ZM domain; the long isoforms possess three C-terminal LIM domains, whereas the short isoforms possess only the PDZ and ZM domains. Zasp/Cypher-knockout mice die within the fifth day after birth (Huang et al. 2003; Zhou et al. 2001). They show muscular weakness, respiratory failure, dilated cardiomyopathy, and difficulty in suckling. Muscle ultrastructure shows discontinuous Z-lines and uncompacted sarcomeres. However, muscle damage seems to occur as a result of contraction. Indeed, whereas cardiac muscle, which begins to contract during embryonic life, shows signs of damage starting from embryonic life, damage to the diaphragm occurs only after birth, when this muscle first starts to contract. Thus, ZASP does not seem to be involved in sarcomerogenesis but instead appears to be involved in Z-line maintenance, possibly acting to reinforce this structure during muscle contraction (Zhou et al. 2001; Faulkner et al. 2001).

In humans, mutations in ZASP are associated with dilated cardiomyopathy, with left ventricular non-compaction (LVNC; Vatta et al. 2003; Xing et al. 2006), and with myofibrillar myopathy (MFM; Selcen and Engel 2005; Griggs et al. 2007). The full spectrum of “zaspopathies” thus encompasses symptoms that are characteristic of MFM, such as Z-line disgregation, and proximal and distal muscle weakness, cardiomyopathies, and peripheral neuropathies (Selcen and Engel 2005).

In order to understand the molecular mechanism(s) involved in the pathogenesis of MFM and cardiomyopathy, suitable animal model systems are needed in which to perform, in depth, studies of the molecular and physiological processes resulting from the dysfunction of the disease gene. In recent years, Drosophila melanogaster has proved to be the choice model on many occasions (Bilen and Bonini 2005; Crowther et al. 2006; Feany and Bender 2000; Wolf et al. 2006; Zordan et al. 2006), as it offers the advantages of extreme versatility, a rich genetic tool box, and a sufficiently simple biological organization. In this respect, a recent paper by Jani and Schock (2007) has reported the isolation of a strongly hypomorphic dzasp deletion mutant that results in recessive early embryonic or first-stage-larval lethality. Furthermore, muscles in such mutant larvae have been shown not to form Z-lines and not to recruit alpha-actinin to the Z-line, leading to the detachment of muscles at the myotendinous junctions. In this paper, we present a molecular characterization of dzasp, an analysis of its larval transcriptional profile, and a functional analysis of the defects resulting from its targeted post-transcriptional silencing. dsRNA interference (dsRNAi) of dzasp has been made possible by the generation of transgenic lines harboring constructs that, through the use of the UAS/Gal4 binary system, have enabled us to drive dsRNAi of dzasp in a tissue-specific manner. Knockdown individuals, produced by using a ubiquitous or a mesoderm-specific Gal4 driver, show pupal lethality thus allowing the functional analysis of larval stages in order to establish the pattern of dzasp expression in muscles and its involvement in muscle structure and function. Our results show that, in addition to the flexibility afforded by the Gal4-driven temporal or spatial modulation of the knockdown, the ubiquitous or mesoderm-specific post-transcriptional silencing of dzasp represents a useful model reproducing some of the key characteristics typical of human zaspopathies.

Materials and methods

Fly strains

Flies were raised on standard yeast–sugar–agar medium (Roberts and Standen 1998) and were maintained at 23°C and 70% relative humidity, under 12-h light/12-h dark cycles. The w1118 strain was microinjected with the transformation plasmid and was used as a control. The dominantly marked, multiply inverted, balancer chromosome w; CyO/Sco; MKRS/TM6B stock was used for determining the chromosomal locations of the transgenes and for subsequent manipulations of the transgenic lines. The following Gal4 lines were used to drive the expression of the UAS–dzasp–IR construct: y, w; Act5C–Gal4/TM6B, Tb (Bloomington Stock Center; genotype: y[1] w[*];Pfw[1mC]¼Act5C–Gal4g17bFO1/TM6B, Tb[1]); How 24B-Gal4 (Brand and Perrimon 1993) and elavC155-Gal4 (Lin and Goodman 1994).

Plasmid construction

The procedure adopted was as detailed by Piccin et al. (2001). In brief, an 880-bp fragment from the coding sequence of the Drosophila gene, dzasp (CG30084), was amplified by using the pair of primers 5′-GAATTCGCTATGCTCGTCCGTTCG-3′ and 5′-CTCGAGGCCTCCACCCATCTGTCG-3′ and cloned in the T vector pDK101 to yield pDK–dzasp. This pair of primers amplifies a sequence that spans exons 14, 16, 19, 21, and 22 (see Fig. 1). A sub-fragment included between the endogenous EcoRI and PstI sites was then excised and cloned in the pBC KS1 vector (Stratagene, La Jolla, Calif.). In parallel, a 330-bp fragment of the green fluorescent protein (GFP)-coding sequence was amplified by using the primers 5′-ACGGCCTGCAGCTTCAGC-3′ and 5′-GAGCTGCAGGCTGCCGTCCT-3′. pDK–dzasp was then linearized and recircularized with the 330-bp PstI–PstI GFP fragment, and the dzasp fragment included between the SalI and PstI sites was excised from pBC. The GFP fragment was included as a heterologous spacer to favor the formation of the double-stranded hairpin, once the dzasp-IR was expressed in the host organism. Finally, the EcoRI–EcoRI fragment containing the two dzasp–IR, separated by the GFP spacer, was cloned into the Drosophila transformation vector pUAST to give UAS–dzasp–IR.
Fig. 1

Diagram showing the exon structure of various transcript isoforms isolated following amplification of larval mRNA via quantitative reverse transcription with polymerase chain reaction (QRT-PCR) and 3′-5′ rapid amplification of cDNA ends. Top Complete structure of the gene to scale (excluding introns). Middle Exons isolated in this study and represented in the isoforms listed below. Bottom left Exon structure of the various isoforms isolated in this study. Font colors: black exons, light blue identified putative 5′ untranslated region (UTR), red identified putative 3′ UTR. Filled colored boxes: red region encoding PDZ domain, light blue region encoding the zasp motif, green regions encoding LIM domains. Bottom right Table showing further details corresponding to each of the isoforms. In particular, in each case, the transcript to the left of each row in the table is referenced, providing the length of the transcript (in kb), the molecular weight of the putative protein (in kDa), and its length in amino-acids (AA). The presence of PDZ, ZM, or LIM domains in the given isoform is indicated (+). In some cases, the transcripts apparently include exons that should encode for a given domain, but the domain does not appear in the putative protein. This occurs because the domain in question is actually encoded only when the corresponding exon is spliced in the proper frame

P-element-mediated transformation

Transformation of Drosophila embryos was carried out according to Rubin and Spradling (1982). Three independent transgenic lines were obtained and named dzasp 67, 73, and 87, two of which carried a single UAS–dzasp–IR autosomal insertion (73dzasp, 87dzasp), and one of which carried two autosomal insertions (67dzasp). Chromosomes harboring the UAS–dzasp–IR insert were balanced and later made homozygous. In situ hybridization was used to map the position and to detect the number of inserts in each line as described in Piccin et al. (2001).

Egg-to-adult viability

For each of the transgenic lines (67, 87, and 73) and for strain w1118, around 300 fertilized eggs were collected on standard yeast–glucose–agar medium (Roberts and Standen 1998) in a Petri dish (60×15 mm), and the same procedure was adopted for the corresponding knockdown individuals in which, in turn, each of Act–Gal4, 24B-Gal4, or elavC155-Gal4 was used to induce dzasp gene silencing. The fertilized eggs were incubated at 23°C, and for each experimental condition, the number of individuals reaching the third instar larva, pupa, or adult, and the relative percentages were calculated. Since the precise staging of large numbers of individuals is difficult during the first larval stages of development, survival was determined collectively for the whole period between egg hatching to the end of larval development (see also Zordan et al. 2006).

RNA extraction and real-time polymerase chain reaction

Total RNA was extracted from approximately 10 larvae with Trizol Reagent (Invitrogen) following the manufacturer’s instructions. mRNA was purified from ~100 µg total RNA with Dynabeads mRNA purification kit as recommended by the manufacturer.

Reverse transcription and real-time polymerase chain reaction

These were performed as in (Zordan et al. 2006) by using the following sets of primers (numbers following the letters “zasp” refer to exon numbers, whereas the letters “f” or “r” refer respectively to “forward” and “reverse”):

In the following lists:

Primers used in real-time polymerase chain reaction (PCR)

zasp13f

CAGTAGCAACGGTAGCACCA

zasp14r

TTGGCCATAAATCGAAGAGG

zasp5f

TACCAGGGCGATCGCTCC

zasp6r

GCGCTGACATGGCGAGTG

zasp1UTRf

CAACTCTCCCGGTCACGGTC

zasp1r

CAGCTGTGGTTGGGCCAT

Rapid amplification of cDNA ends and polymerase chain reaction

The expression of dzasp isoforms was investigated by reverse transcription (RT)-polymerase chain reaction (PCR) from larval mRNA. The 5′ and 3′ rapid amplification of cDNA ends (RACE) was performed by using the Smart Race cDNA amplification kit according to the manufacturer’s instructions; 1 µg mRNA was used for first-strand cDNA synthesis.

PCRs were carried out using the following primer pairs (“gsp” indicates gene-specific, and “ngsp” indicates non-gene-specific, both as in the Smart RACE kit intructions):

Primers used in 5′-3′ rapid amplification of cDNA ends

zasp13gsp1

AGCTGGGCTATCTGTTGCAGTTGCTGC

zasp13ngsp1

CAGCGGATGGTGCTACCGTTGCTACTG

zasp20gsp1

GGCGTGGCAAGGTGGCGAAGGTTTGAGG

zasp20ngsp1

GGCAGCGCTTTGAGCTCCTGTGGGC

zaspr2UTR

GTGGTGCACTGCCAGTTCAGTTCTTGGG

zasp2r3UTR

CAGTTCTTGGGCGACGTTAACTAAGCGACG

zasp13gsp2

CAGTAGCAACGGTAGCACCATCCGCTG

zasp13ngsp2

GCAGCAACTGCAACAGATAGCCCAGCT

zasp5gsp2

GGAATACCAGGGCGATCGCTCCGAGG

zasp5ngsp2

GCGCGAGGAGGAGACCGGCCAGTCC

PCRs were performed in a 50-µl reaction mixture containing 1× Expand High Fidelity Buffer, 1.5 mM MgCl2, 10 pmol each primer, 200 µM each dNTP, 50 ng cDNA, 2.6 U Expand High Fidelity enzyme mix (Roche). Following a 2-min denaturation step at 94°C, 35 rounds of amplification were performed (94°C for 30s, 60°C–68°C for 30 s, 72°C for 1–4 min, plus a 5-s cycle elongation for each successive cycle). This was followed by incubation at 72°C for 10 min. The PCR products were analyzed by 1% agarose gel electrophoresis. All RT-PCR products were cloned in pCR2.1-TOPO (Invitrogen) and were sequenced to confirm their identity.

Full-length amplifications were carried out using the following primers (numbers following the letters “zasp” refer to exon numbers, whereas the letters “f” or “r” refer respectively to “forward” and “reverse”):

Primers used for full-length PCR

zasp2UTRf

AGTTGAGAGAAAATCGTCGCTTAG

zasp24r

TTAGCGCGCGTGATTCTTGCAG

zasp4UTRf

AGTTTCTTTGCCCCCTGTGCTC

zaspl7r

AAGTTTATATTCGCAGTTTATTGC

zasp1UTRf

CAACTCTCCCGGTCACGGTC

Anti-dZASP antibody generation

A polyclonal antibody specific for a known region of Drosophila ZASP was produced by inoculating 25 µg specific peptides resuspended in phosphate-buffered saline (100 µl) intraperitoneally in BalbC female mice. These mice were immunized from 2 months of age onwards every 21 days. Blood samples were taken 7 days after inoculation, and sera were produced. The blood was taken from the saphenous vein, and the mice could be bled in this way every 3 weeks without trauma. Injections were given by using Freund’s incomplete adjuvant. The peptide used was conjugated to bovine serum albumin (BSA) at the C-terminal, the sequence of the peptide being: GATSAPKRGRGILNKC–BSA-conjugated (encoded by exon 19).

Protein extraction and SDS-polyacrylamide gel electrophoresis

Approximately 90 mg (corresponding to 10 larval body walls, dissected as reported above) were homogenized in 200 μl lysis buffer (20 mM HEPES pH 7.5, 100 mM KCl, 5% glycerol, 10 mM EDTA, 1% Triton X-100) with a protease inhibitor cocktail (1 mM dithiothreitol, 0.5 mM phenylmethane sulfonyl-fluoride, 20 μg/ml aprotinin, 10 μg/ml leupeptin, 5 μg/ml pepstatin A). The solution was spun at 12,000g for 10 min, and the precipitate was discarded. The supernatant was spun again at 12,000g for 3 min. Protein quantification was conducted by using the Bradford assay (Bio-Rad), and equal amounts of total proteins for each sample were resolved on a precast 3%-8% TRIS-acetate polyacrylamide gel (NuPAGE, Invitrogen). Following separation and Western blotting onto nitrocellulose membrane (Trans-Blot Transfer Medium, Bio-Rad), the membrane was blocked for 1 h in 0.01 M TRIS-HCl pH 7.5, 0.14 M NaCl, 0.1% Tween 20 with 5% dry milk (Carnation NonFat Dry Milk) and incubated overnight at 4°C with mouse anti-dZASP antibody (1:200). An anti-mouse IgG conjugated to horseradish peroxidase (1:5000; Bio-Rad) was used as the secondary antibody. Positive immunoreactivity was visualized by using a chemiluminescent system. The immunological specificity of the anti-dZASP antibody was confirmed by performing a peptide/antigen competition test for the antibody. The peptide concentration was determined by an enzyme-linked immunosorbent assay.

Anti-dZASP staining

Third instar larvae were dissected and treated as described in Beramendi et al. (2005) with little modification. Larvae were fixed for 30 min in 4% paraformaldehyde (PFA) and incubated overnight with anti-dZASP antibody at a 1:300 dilution. Secondary antibodies were tetramethylrhodamine-isothiocyanate-conjugated rabbit anti-mouse IgG (DAKO; 1:500 dilution for 2 h) or fluorescein-iosthiocyanate-conjugated goat anti-mouse IgG (Sigma; 1:500 dilution) when used for double-staining together with phalloidin-rhodamine).

Phalloidin-rhodamine staining

Third instar larvae were dissected and treated as described in Beramendi et al. (2005). Samples were fixed for 15 min in 4% PFA, incubated for 20 min in phalloidin-rhodamine (Sigma; 1:1000) and then observed with a fluorescence confocal microscope (Radiance 2000; Bio-Rad, mounted on a Nikon Eclipse E600 microscope), under a 40× Nikon Plan Fluor N.A.=0.75, DIC M objective. Measurements were made on digitized images by using ImageJ software. In control and knockdown larvae, four parameters relating to muscle fiber 6 were measured: length (l), width of the two extremities, viz., the head end (he) and tail end (te), where the ends were defined by the extremities of the muscle fiber attached to the tendons (with the head end being toward the anterior and the tail end toward the posterior), and width of the central region of the fiber (c). These parameters were used to calculate the visible area of muscle fiber as follows: \( \left( {\left( {\left( {{\text{he}} + {\text{c}}} \right)*{\text{l}}/2} \right)/2} \right) + \left( {\left( {\left( {{\text{te}} + {\text{c}}} \right)*{\text{l}}/2} \right)/2} \right) \), considering the muscle as the geometrical equivalent of two juxtaposed trapeziods (the central part of the fiber tending to be thinner than the extremities). Ten larvae per genotype and two fibers per larva were measured.

All experiments were performed on individuals of the three transgenic lines. Most of the data presented refer to transgenic line 67, since this line showed the lowest amount of residual mRNA (i.e., the strongest level of zasp mRNA knockdown).

Electron microscopy

Transmission electron microscopy of muscles from larval body wall preparations was performed essentially as described in Zordan et al. (2006). Briefly, third instar larvae were dissected to obtain larval body wall fillets. These were then transferred to ice-cold fixation solution and fixed for 6 h. The next day, larvae were washed and treated for 2 h with post-fixative solution, rinsed, and then dehydrated through an ethanol series, washed in acetone, and finally embedded in resin. Ultrathin cross sections of larval body wall muscles were stained in uranyl acetate solution followed by lead citrate and finally rinsed. Sections were examined and photographed with a Philips 200 electron microscope.

Locomotion

Larval locomotion was tested with two assays on individual third instar larvae of similar length placed in the center of a 9-cm Petri dish coated with a thin layer of non-nutritive agar. In the “roll-over” assay, the larva was turned upside down with a fine brush, and the time taken for the larva to turn back to its normal position (ventral side down) was measured. Ten larvae were tested for each genotype, and the test was performed ten times for each larva. Each larva was thus assigned a value corresponding to the average of the ten measurements. Larval locomotor speed was determined by the “crossover” test: third instar larvae (20 for each genotype) were analyzed in 100×15-mm Petri dishes, with 1% agar covering the base to a depth of 0.5 cm. Individual larvae were placed into a narrow linear groove produced with the aid of a glass capillary (diameter 1 mm) on the surface of the agar layer and allowed to acclimatize for 60 s. Testing was performed by using a fiber optic lamp (100 W) with a photon throughput of 60 μmol/m2 per second (light intensity was measured by a Basic Quantum Meter, model QMSW-SS; Apogee Instruments). The time necessary to cover a 1-cm distance was recorded, and larval locomotor speed was expressed in centimeters per second. Each larva was tested three times, and the resulting data were expressed as averages. Statistical analyses were performed by using Student’s t-test.

Results

Flybase reports six transcript isoforms for CG30084: named CG30084-RG, -RA, -RE, -RF, -RC, and -RH. Isoforms RG, RA, RE and RC encode for putative proteins having the following domains (proceeding from N-terminus to C terminus): PDZ-ZM-LIM---------(LIM)3.

RF encodes a putative protein with a PDZ-ZM-LIM structure. RH encodes for a putative protein lacking the N-terminal PDZ, ZM and LIM domains and having only the triple C-terminal LIM domains.

A search with SMART (http://smart.embl-heidelberg.de/smart/) employing, as a query sequence, either one of the long Drosophila isoforms reported in Flybase (RG, RA, RE, or RC) has revealed that, among mammals, known proteins with a similar domain architecture show a PDZ-ZM---------(LIM)3 organization. An extra LIM domain immediately adjacent to the ZM domain appears to be common to insects (Drosophila spp., Anopheles spp., Apis mellifera, and Tribolium castaneum) and to Caenorhabditis spp. Indeed, this organization is probably ancestral to that found in mammals according to Te Velthuis et al. (2007) who postulate that the PDZ-ZM---------(LIM)3 organization may have evolved from the PDZ-ZM-LIM---------(LIM)3 type following the loss of the N-terminal LIM domain.

By virtue of the presence of all the functional domains present in the mammalian ZASP protein (with particular reference to the N-terminal PDZ and ZM domains and the C-terminal triple LIM domains), we consider the putative protein encoded by CG30084 to be the Drosophila ortholog of mammalian ZASP (see also Jani and Schock 2007). In particular, the N-terminal PDZ-ZM region shows a 31% identity and a 52% similarity with the corresponding human ZASP domains, whereas the C-terminal (-LIM3) domains show a 47% identity and a 61% similarity with the corresponding human ZASP domains.

Expression studies

The analysis of transcripts established by a combination of quantitative RT-PCR and 5′ and 3′ RACE is shown in Fig. 1. In particular, since the silencing of dzasp in transgenic line 67 leads to 100% pupal lethality, we have characterized the transcriptional profile of dzasp at the larval developmental stage. The results are complex, suggesting that the transcription of this locus gives rise to multiple splice variants. This situation is not unlike that found in mammals in which multiple splice variants have also been isolated (Faulkner et al. 1999; Huang et al. 2003). Although a comparison of larval and adult transcripts is not straightforward, it is nonetheless useful to compare the results published by other authors (Machuca-Tzili et al. 2006; Jani and Schock 2007) with our own and with those present in Flybase. For this purpose, we have chosen to number the transcripts described in this paper consistently with the numbering scheme adopted by Jani and Schock (2007), as shown in Fig. 1. In addition, Table 1 allows a direct comparison between exons as reported by us, Jani and Schock (2007), Machuca-Tzili et al. (2006) and Flybase.
Table 1

Synopsis of dzasp exons as identified in the present paper, by Flybase and in published reports by Machuca-Tzili et al. (2006) and Jani and Schock (2007)

Genomic landmark

Exon length (bp)

Flybase

Machuca-Tzili et al. (2006)

Jani and Schock (2007)

This papera

Domain encoded

Start

Stop

11704509

11704265

244

partial 27

  

5′UTR-1

 

11702419

11702234

185

26

1

1

5′UTR-2

 

11702402

11702234

168

25

  

5′UTR-3

 

11690993

11690864

129

24

2

2

1

PDZ

11670973

11670820

153

23

3

3

2

PDZ

11670750

11670593

157

22

4

4

3

 

11669411

11669278

133

21

5

5

4

ZM

11668585

11668490

95

20

6

6

5

 

11667785

11667563

222

19

7

7

6

 

11667166

11666770

396

18

8

8

7

 

11666051

11665857

194

17

9

9

8

LIM

11664791

11664681

110

16

10

10

9

 

11663542

11663278

264

partial 15

  

5′UTR-4

 

11663013

11662815

198

14

11a

11

10

 

11663012

11661933

1079

13

11abc

 

11

 

11662598

11661925

673

12?

11ac

12

12

 

11662263

11661933

330

11

11c

 

13

 

11661472

11661319

153

10

12b

13

14

 

11661471

11661285

186

9

12ab

 

15

 

11660760

11660544

216

8

 

14

16

 

11658654

11658459

195

?

13

15

3′UTR-17

 

11657056

11652628

4428

7

14abc

 

18

 

11652759

11652628

131

6

14c

16

19

(LIM)

11652342

11651431

911

5

15

17

20

(LIM)

11651337

11651150

187

4

16

18

21

LIM

11651096

11650855

241

3

17

19

22

LIM

11650775

11650698

77

2

  

23

 

11650776

11650063

713

1

18

20

24

LIM

aUTR Untranslated region

Production of dzasp knockdown and quantification of residual mRNA

We generated three independent Drosophila lines bearing one or more insertions of a construct designed to produce post-transcriptional silencing of dzasp. In what follows, for each line, the insert map position (chromosome number followed by the cytogenetic band position) is given in parentheses: line 67 carries two insertions (2R 57A/B and 3L 75A/B); line 73 carries one insertion (3L 66A); line 87 carries one insertion (3L 61D). Post-transcriptional gene silencing of dzasp via dsRNAi, was obtained by crossing the UAS-dzasp-IR lines to driver lines expressing tissue-specific Gal4. The driver lines employed were: How24B-Gal4 (24B-Gal4; mesodermal derivatives and muscular tissue specific), Actin5C-Gal4 (ubiquitous), and elav-Gal4 (panneuronal).

The Actin5C (Act–Gal4) driver induces early ubiquitous expression of dsRNAi-mediated knockdown. Table 2 shows the residual amount of dzasp mRNA determined by real-time PCR following knockdown.
Table 2

Percentage (as compared to controls) of residual dzasp mRNA in ubiquitous knockdown (KD) individuals. In each case, the control was the respective parental dsRNA-bearing transgenic line. RT-PCRs were carried out in triplicate for each genotype

Transgenic Line

Percentage residual mRNA

67 dzasp-Act-Gal4-KD

17%

73 dzasp-Act-Gal4-KD

42%

87 dzasp-Act-Gal4-KD

33%

The Western blot analysis of larval body wall extracts is shown in Fig. 2. The pattern of bands obtained by using anti-ZASP antibody on wild-type larval extracts can be seen in lane 1, whereas lanes 2–4 show the corresponding patterns obtained with extracts from each of the Act-Gal4 knockdown lines (67, 73, and 87, respectively). Two bands (of molecular weight approximately 70 and 80 kDa) are clearly missing from lanes 2–4, suggesting that the antibody recognizes two isoforms that are undetectable in the knockdown lines. The protein isoform(s) producing the 80-kDa band might correspond to the putative proteins encoded by transcripts 6 and 7 (see Fig. 1), whereas the 70-kDa band is most closely approximated by the putative product of transcript 2 (predicted weight: 63 kDa; Fig. 1), although other splice variants that we may have not isolated might nevertheless exist.
Fig. 2

Western blot of larval body wall protein extracts. Lanes from left to right are extracts from: lane 1 w1118 controls, lane 2 24B-Gal4 knockdown larvae from line 67, lane 3 24B-Gal4 knockdown larvae from line 73, lane 4 24B-Gal4 knockdown larvae from line 87. Knockdown (KD) individuals (lanes 2–4) lack two bands of approximate molecular weight (from top to bottom) of 80 kDa and 70 kDa, respectively; these are clearly present in the w1118 controls

Developmental effects of dzasp knockdown

We determined the percentage of individuals that survived to various developmental stages. The most dramatic effect caused by the lack of dzasp was seen when dsRNAi was driven ubiquitously, resulting in 100% pupal lethality (Table 3). Furthermore, in the case of the ubiquitous knockdown of dzasp, the percentage of embryos that developed to third larval instar was strongly reduced compared with controls. Nonetheless, practically all surviving 3rd instar larvae developed to the pupal stage, regardless of the tissue specificity of dsRNAi. The lack of dZASP thus appeared to be particularly critical for the larval-to-adult metamorphosis. This effect was maximal (100% lethality) when dsRNAi was driven ubiquitously but remained evident (53%) even when a muscle-specific driver was employed. A weak effect was observed in the survival of embryo to 3rd instar larva when a panneuronal driver was employed, whereas the progression through the successive stages of development did not appear to be sensitive to the lack of dZASP at the nervous system level (Table 3).
Table 3

Survival rates from embryo to 3rd instar larva, from 3rd instar larva to pupa, and from pupa to adult stage of development of individuals in which dzasp expression was knocked down by using a ubiquitous driver (Act-Gal4), a mesodermal specific driver (24B-Gal4), or a panneuronal driver (elav-Gal4). Survival rates for each genotype were determined starting from 300 eggs each. Numbers represent the percentage of the number of individuals that survived for each stage with respect to concurrent controls. In each case, the control was the respective parental dsRNA-bearing transgenic line (KD knockdown)

Transgenic line

Embryo to 3rd instar larva

3rd instar larva to pupa

Pupa to adult

67 dzasp-Act-Gal4-KD

66%

100%

0%

67 dzasp-24B-Gal4-KD

100%

97%

53%

67 dzasp-elav-Gal4-KD

87%

99%

100%

dZADP localizes at the Z-lines in body wall muscles of larval D. melanogaster

Given the homology with mammalian ZASP, we investigated the expression and subcellular localization of dZASP in the muscle tissue of larval Drosophila. To this end, we evaluated the immunofluorescence staining of larval body wall muscles by using our mouse-anti-dZASP antibody. Double-staining of larval body wall preparations with the anti-dZASP antibody and with phalloidin-rhodamine revealed that the anti-dZASP narrow bands were localized in a central position within the wider phalloidin-positive bands (actin filaments; Fig. 3), suggesting that dZASP localized to the Z-line analogously to the mammalian homolog (Faulkner et al. 1999; Zhou et al. 1999). This was also in agreement with the co-localization of dZASP and alpha-actinin shown for Drosophila larval muscles by Jani and Schock (2007). The above analysis was conducted on larval muscle 6. Drosophila larval muscles 6 and 7 are among the largest muscles present in the larva. They are present in a ventral and longitudinal position and have been used extensively for detailed molecular and electrophysiological studies by many researchers in the field, one of the advantages also being that they are innervated almost exclusively by glutamatergic-type synapses (Guan et al. 1996). From the purely muscular point of view, these muscles are representative of all the muscles present in the larval body wall and viscera, and all such muscles are of the supercontractile type, the peculiar characteristic of supercontractile muscles being their capacity to contract largely below 50% of their resting length. The muscle fibers analyzed in our studies were always fibers 6 and/or 7 from right or left larval hemi-segments 2-4.
Fig. 3

Confocal image of muscle fibre 6 from the body wall of a w1118 larva. Staining was performed with both phalloidin-rodhamine (red) and anti-dZASP antibody (green). Anti-dZASP staining reveals a pattern of narrow stripes located centrally within the band stained by phalloidin-rhodamine and corresponding to the Z-lines. Bar 5 μm

Effects of dzasp knockdown on larval locomotion

Since dZASP was expressed in muscle and localized to the Z-lines, we hypothesized it might have a role in muscle contraction, and that a lack of dZASP at this level might affect larval locomotion. We measured the time that a larva required to travel a distance of 1 cm. In parallel, we also conducted a “roll-over” test, which measured the time taken for a larva placed in a ventral-side-up position to roll-over into the normal ventral-side-down position. dzasp knockdown by using a muscle-specific driver (24B-Gal4) led to a significant decrease in the speed of locomotion and to a poorer performance in the roll-over test compared with controls (Fig. 4a, b); this decrease was correlated with a strong reduction of the dZASP signal (Fig. 5; see also below). On the contrary, when dzasp expression was silenced ubiquitously, the roll-over performance appeared to be influenced more dramatically than the locomotor speed. No locomotion impairments were detected when dzasp expression was knocked down by using the panneuronal driver (elav-Gal4; Fig. 4a, b).
Fig. 4

a Larval locomotion test. Mean times (error bars standard error) taken for larvae to travel 1 cm. 24B-Gal4 knockdown larvae (24B-KD) are significantly slower than controls (w1118), but Act-Gal4 knockdown larvae (Act-KD) only show a weak reduction in locomotor speed. *P<0.05. Data are from 20 larvae for each genotype. b Roll-over test. Mean time (error bars standard error) taken for larvae to return to the ventral-side-down position. 24B-Gal4 knockdown larvae (24B-KD) and Act-Gal4 knockdown larvae (Act-KD) are significantly slower than controls (w1118), although Act-Gal4 knockdown larvae show a much stronger phenotype. *P<0.05. Data are from 10 larvae for each genotype

Fig. 5

Confocal images showing muscle fibre 6 stained with an anti-dZASP antibody. Interference is driven in muscle tissue only (24B-Gal4). The control (a, w1118) shows a regularly repeating striated pattern. In the knockdown lines (b24B-Gal4 knockdown larvae from line 67, c24B-Gal4 knockdown larvae from line 73, d24B-Gal4 knockdown larvae from line 87), fluorescence is reduced in agreement with the relative amount of residual mRNA determined by QRT-PCR (Table 2). Bar 10 μm

Post-transcriptional silencing of dzasp leads to a dramatic reduction in protein expression in larval muscular tissue

Following dzasp silencing, the anti-dZASP antibody was used to detect the absence of dZASP in larval muscles. The larval body wall preparations of transgenic knockdown larvae were stained with an anti-dZASP antibody showing a reduction in the fluorescence intensity in muscles of knockdown larvae compared with the controls (Figs. 6, 7). The level of reduction seen by fluorescence was in keeping with the reduction in dzasp mRNA following knockdown (Table 2). Indeed, the lowest fluorescence intensity was detected in larvae from transgenic line 67, which carried two copies of the dsRNAi construct, and which presented the lowest amount of residual mRNA following knockdown (17%, see Table 2). Furthermore, when the knockdown was driven in mesodermal derivatives by using the 24B-Gal4 driver (i.e., mainly muscle tissue), a stronger reduction of the dZASP signal (Fig. 5) was obtained. Nonetheless, at this level of observation, no morphological defects were visible in muscle structure.
Fig. 6

Confocal images showing muscle fibre 6 stained with anti-dZASP antibody. Interference is driven ubiquitously (Act5C-Gal4). The control (a, w1118) shows a regularly repeating striated pattern. In the Act-Gal4 knockdown lines (bAct-Gal4 knockdown larvae from line 67, cAct-Gal4 knockdown larvae from line 73, dAct-Gal4 knockdown larvae from line 87), fluorescence reduction is much less evident than in the case of 24B-Gal4 knockdown individuals (Fig. 5), although the slight reduction is in agreement with the relative amount of residual mRNA determined by QRT-PCR (Table 2). Bar 10 μm

Fig. 7

Electron-microscopy images showing the structure of a transverse section of body wall muscle from (a) wild-type control larva, (b) Act-Gal4 knockdown larva, and (c) 24B-Gal4 knockdown larva. Higher magnification images of boxed areas in b, c are shown in d, e, respectively. Cross sections of larval muscles 6 or 7. Muscle fiber structure is clearly degenerated in the knockdown individuals. In particular, degeneration (see vacuolizations in b–e) is more severe in the case of 24B-Gal4 knockdown larval muscles (c) than in Act-Gal4 knockdown larval muscles (b). Bars 2 μm (a–c), 0.54 μm (d, e)

Electron microscopy

Electron-microscopic analysis of larval muscle fibers 6 and 7 (Fig. 7) showed that the silencing of dzasp with either the Act-Gal4 or the 24B-Gal4 driver produced significant alterations in muscle fiber substructure. In particular, transverse sections of muscle fibers from a dzasp (Act-Gal4) knockdown larva (Fig. 7b, d) and a dzasp (24B-Gal4) knockdown larva (Fig. 7c, e) showed that the myofibrils were less regularly and less densely packed than those of the corresponding control (Fig. 7a). Furthermore, the extent of such morphological alterations appeared to be stronger in the case of the 24B-Gal4 knockdown individuals (Fig. 7c, e) than in the Act-Gal4 knockdown samples (Fig. 7b, d).

Ubiquitous dzasp silencing causes muscle hypertrophy

We analyzed the possible effects of dzasp knockdown on muscle fiber growth. We noticed that, when dzasp was silenced in all tissues (i.e., when using the Act5C-Gal4 driver), muscle fiber 6 area was significantly larger than that in controls (Fig. 8), whereas muscle fiber 6 area in dzasp (24B-Gal4) knockdowns was not altered with respect to controls.
Fig. 8

Visible area of muscle fibre 6. Ubiquitously driven (Act-Gal4-driven) dzasp knockdown leads to muscular hypertrophy, whereas 24B-Gal4 knockdown does not influence muscle size. Data represent the mean (error bars standard error) of 20 muscle fibers from ten larvae for each genotype analyzed. *P<0.05

Discussion

The mammalian ZASP protein is highly expressed in skeletal and cardiac muscle and co-localizes with alpha-actinin at the Z-line, possibly functioning in the stabilization of sarcomere structure during contraction and with a supposed function in signaling (Faulkner et al. 1999; Zhou et al. 1999). In the present study, the anti-dZASP staining of Drosophila larval preparations has revealed that the protein is expressed in larval muscle tissue and localizes to the Z-line. This pattern is reminiscent of the localization of mammalian ZASP. In mammals, ZASP binds to alpha-actinin-2 at the Z-line through its PDZ and ZM domains. These domains are conserved in dZASP, and thus, binding to the Z-line in Drosophila probably occurs similarly (see also Jani and Schock 2007). In order to investigate the functional implications deriving from a lack of dZASP, we have employed dsRNAi in order to produce post-transcriptional silencing of the gene. Furthermore, in order to explore the consequences deriving from a lack of dZASP in the various regions of the organism, we have knocked down gene expression by driving dsRNAi either ubiquitously, in mesodermal derivatives, or in the nervous system.

In mammals, no direct evidence has been presented that ZASP has a function in tissues other than muscles. Nevertheless, several papers report that the protein is expressed at low levels in other tissues (Faulkner et al. 1999; Zhou et al. 1999; Arimura et al. 2004). In addition, some patients affected by myopathies consequent to mutations in the ZASP gene, exhibit symptoms that apparently do not directly involve muscle tissue, e.g., in the ZASP-dependent neuropathies (Selcen and Engel 2005). Our results show that, following the knockdown of dzasp in Drosophila, a lack of protein in the whole organism has a dramatic effect on development. In particular, it almost halves the percentage of embryos that reach the 3rd larval instar. The effect is more dramatic when dZASP is absent during metamorphosis, when 100% of individuals die before reaching the adult stage. The absence of the protein only in mesodermal derivatives affects the survival rate up to metamorphosis to a lesser extent (53%, see Table 3). Taken together, these results suggest that dZASP is important in muscle tissue during metamorphosis, but that its role, at this developmental stage, appears not to be restricted to this district alone, as evidenced by the much stronger developmental effect obtained by driving the knockdown of dzasp ubiquitously. On the other hand, a lack of dZASP at the nervous system level, obtained by driving the knockdown with the elav-Gal4 driver, results in mild mortality restricted to the embryonic stages of development. In this respect, Jani and Schock (2007) have recently reported a Drosophila hypomorphic mutant for dzasp that lacks most of the dzasp gene products giving a more severe phenotype with mortality in the early stages of larval development. A comparison of the exons removed by the deletion in the mutant generated by Jani and Schock (2007) and the exon structure of the isoforms isolated in this study shows that their mutant probably has a deletion of exons 4-8 (see Fig. 1); this implies that at least two isoforms (i.e., isoforms 5 and 10, Fig. 1) characterized in the present study might still be expressed in the mutant of Jani and Schock (2007). Indeed, similar isoforms have also been described by these authors as supplementary data to their report (isoforms HL08122 and GH15137), and one of these (HL08122) corresponds exactly to isoform 10 (Fig. 1). In addition, the finding that such isoforms can still be encoded by the deletion mutant has led to the realization that this mutant might be a hypomorph and not a complete amorph (Jani and Schock 2007). Interestingly, these isoforms do not encode for the N-terminal PDZ, ZM, and LIM domains, but only for the C-terminal triple LIM domains. The role of such isoforms remains, for the time being, unknown. Importantly, these isoforms would be targeted for knockdown in our model, since the only isoform (among those isolated by us) that is possibly not subject to knockdown is isoform 8 (Fig. 1), as this isoform does not include any of the exons that constitute the dsRNA used to produce interference. This in turn suggests that our model and that of Jani and Schock (2007) can be viewed as complementary, given the partial overlap of the exons knocked out or knocked down, as might be the case, in each of the two studies. In our knockdown individuals, death at the pupal stages of development might be the consequence of the lack of protein at the muscle level; this, during metamorphosis, might result in a malfunctioning of the muscles, the contraction of which is required during pupal eclosion (Fristrom and Fristrom 1993). Indeed, the lack of dZASP in larval muscles has a negative effect on both linear and coordinated (i.e., roll-over test) locomotion. Surprisingly, larvae with ubiquitously silenced dzasp show a dramatic deficit in coordinated locomotion, but the effect on linear locomotion is much more limited. In this respect, the muscles of ubiquitously knocked down individuals (i.e., Act-Gal4) show immunocytochemically detectable amounts of residual protein (Fig. 6), whereas the muscles of 24B-Gal4 knockdown individuals show little or no residual anti-dZASP-related fluorescence (Fig. 5). This implies that the lack of ZASP at the muscular level alone (i.e., following 24B-Gal4 knockdown) is not sufficient by itself to result in the roll-over deficiency, which is however significantly affected in the Act-Gal4 knockdown individuals. Contrariwise, Act-Gal4 knockdown individuals show a weaker impairment in the locomotor test, because they present less reduction in ZASP at the muscular level, but the ubiquitous reduction of ZASP (i.e., muscle tissue plus nervous system) results in a significant defect in the roll-over test. Similarly, the knockdown of ZASP in the nervous system alone is not suffcient to produce any of the behavioral defects; this could be interpreted as reflecting the requirement of either a strong deficit of ZASP in muscle tissue (to produce a significant reduction in locomotor speed) or a deficit of ZASP in both muscle and nervous system (to produce a significant impairment in roll-over performance).

The hypertrophy observed in the muscles of ubiquitous dzasp knockdown larvae, but not in 24B-Gal4 (i.e. muscle-specific) knockdown individuals (Fig. 8), suggests that this phenomenon depends upon the concomitant lack of dzasp in muscles and other tissues (as occurs in the Act-Gal4 knockdown larvae). It is worth recalling that 24B-Gal4-driven knockdown results in a stronger reduction in ZASP in muscle as compared with Act-Gal4-driven knockdown. The latter observation could be taken to mean that the hypertrophy observed following Act-Gal4 knockdown reflects the attempt of muscle tissue to compensate for the lack of ZASP, whereas the greater extent of damage occurring following 24B-Gal4-driven knockdown might be beyond any compensation by the muscles. Whatever the mechanism, this hypertrophic phenotype recalls the hypertrophy of the cardiac left ventricular associated with LVNC observed in human subjects carrying mutant forms of ZASP. In Drosophila and man, the defect is possibly consequent to muscle activity, and at least in the case of LVNC, the muscles that are more strongly affected are those subjected to the most exertion, as occurs in the mouse Cypher/Zasp knockout model, in which cardiac and diaphragm muscle damage occurs only after these muscles begin to function (Zhou et al. 1999). Indeed, dzasp embryonic hypomorphs (Jani and Schock 2007) show a muscle detachment phenotype at myotendinous junctions, and this phenotype becomes manifest in late stage 17 embryos, when muscle contractions have begun to occur. In addition, the muscle morphology appears to be more severely altered in the hypermorphs than is the case in our model system. Nonetheless, the observation that, at the electron-microscope level, the dzasp mutant (Jani and Schock 2007) shows disorganized muscle fiber filaments, which appear to lose their parallel array arrangement, is in keeping with the loss of the compaction phenotype observed by us at a similar level of analysis. Strikingly, this loss of compaction is reminiscent of the non-compaction phenotype observed in human LVNC.

Together, the results obtained in this study suggest that the post-transcriptional silencing of dzasp produces a spectrum of features in Drosophila recapitulating some of the key aspects typical of human zaspopathies. This model system, in which the dsRNAi can be modulated in a tissue-specific and/or temporally controlled manner, provides an important complement to the existing hypomorphic dzasp mutant (Jani and Schock 2007). Additional investigative routes can thus be opened with the objective of gaining further insights into the molecular mechanism(s) involved in determining the role of ZASP in the pathogenesis of human muscle disease.

Notes

Acknowledgements

Thanks are due to Mr. M. Sturnega, ICGEB, Trieste, for excellent technical assistance in the immunization of the animals used for antibody production.

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Copyright information

© Springer-Verlag 2009

Authors and Affiliations

  • Clara Benna
    • 1
  • Samantha Peron
    • 2
  • Giorgia Rizzo
    • 1
  • Georgine Faulkner
    • 3
  • Aram Megighian
    • 2
  • Giuliana Perini
    • 2
  • Giuseppe Tognon
    • 4
  • Giorgio Valle
    • 1
  • Carlo Reggiani
    • 2
  • Rodolfo Costa
    • 1
  • Mauro A. Zordan
    • 1
  1. 1.Department of BiologyUniversity of PadovaPadovaItaly
  2. 2.Department of Human Anatomy and Physiology, Section of PhysiologyUniversity of PadovaPadovaItaly
  3. 3.International Centre for Genetic Engineering and BiotechnologyTriesteItaly
  4. 4.CNR Institute of Biomedical TechnologyUniversity of PadovaPadovaItaly

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