© 2002 Wiley-Liss, Inc.
genesis 32:27–31 (2002) DOI 10.1002/gene.10025
TECHNOLOGY REPORT
High-Efficiency Gene Transfer into Adult Fish: A New Tool to Study Fin Regeneration Marcel Tawk,1 David Tuil,2 Yvan Torrente,3 Sophie Vriz,4 and Denise Paulin1 1
Laboratoire de biologie mole´culaire de la diffe´renciation, Universite´ Paris 7, Paris, France ICGM, De´partement de GDPM, Paris, France 3 Dino Ferrari Center, Institute of Clinical Neurology, University of Milan, Milan, Italy 4 UFR de biologie, Universite´ Denis Diderot, Paris, France 2
Received 30 July 2001; Accepted 19 November 2001
Summary: Zebrafish represents an excellent model to study the function of vertebrate genes (e.g., well-developed genetics, large number of mutants, and genomic sequencing in progress), inasmuch as we have tools to manipulate gene expression. Recent use of injected morpholinos in eggs provides a good method to “ knockdown ” gene expression in early development (Nasevicius and Ekker, 2000), and the “caged” RNA injected in eggs allows to overexpress a gene in a specific set of cells (Ando et al., 2001). However, a method to specifically modify gene expression in the juvenile or in the adult is still missing. Such a method would be a very powerful tool to understand gene function in differentiated tissues. We describe here an electroporationbased approach, which allows gene transfer in adult tissues. Its efficiency was assessed using a GFP (green fluorescent protein) dependent assay. We then used this method to disrupt the Fgf signalling pathway during the process of regeneration. genesis 32:27–31, 2002. © 2002 Wiley-Liss, Inc.
Key words: electroporation; fin; zebrafish; Fgf receptor 1; regeneration; dominant-negative mutation; Fgf
Regeneration of appendages represents a very efficient tool to study stem cell recruitment, proliferation and differentiation. The ability to regenerate appears to be widespread throughout metazoans, but in adult vertebrates, the regenerative ability of complex body structures seems to be restricted to Urodeles and Actinopterygiens (Brockes, 1997). Caudal fin regeneration in the teleost Danio rerio, or zebrafish, represents an excellent model to understand epimorphic regeneration (Poss et al., 2000). After amputation, a complete segmented fin regenerates in less than 3 weeks. This process goes through the following steps: (1) formation of a multilayer wound epidermis; (2) recruitment of stem cells and dedifferentiation of mesenchymal cells; (3) proliferation of these cells to form the regenerative blastema; and (4) differentiation of the proximal blastema cells and morphogenesis of the new fin (Johnson and Weston, 1995; Santamaria and Becerra, 1991). Among molecules that
are involved in the growth of the regenerate or that may play a key role in fin patterning are retinoids and their receptors (White et al., 1994), segment polarity genes, Fgfs and their receptors, bone morphogenetic proteins (BMPs), and homeobox containing genes (Geraudie and Ferretti, 1998). However, the respective roles of these molecules during fin regeneration has not yet been ascertained. Functional studies in zebrafish, as in other animal models, call for the ability to manipulate gene expression and to analyze the resulting phenotype. In 1997, Jun-Ichi Okamura group demonstrated that electroporation provides an efficient approach to DNA transfection of somatic cells in live chicken embryos (Muramatsu et al., 1997). This method was rapidly applied to living embryos of other species, such as mice (Akamatsu et al., 1999) or xenopus (Eide et al., 2000), and was used to transfect DNA into various adult tissues such as solid tumors (Goto et al., 2000) or skeletal muscle (Aihara and Miyazaki, 1998; Mir et al., 1999; Swartz et al., 2001). We report here the electroporation conditions to efficiently transfer DNA into adult zebrafish fin cells. We used this method to electroporate a DNA construct encoding a dominant-negative form of the Fgf receptor 1, which disrupted fin regeneration. To observe the expression pattern of the in vivo electrotransferred gene, we injected pCMV-green fluorescent protein (GFP) plasmid DNA (15 g in 15 l) into the dermal skeleton of the fin and then administered 10 15-V pulses with a 60 ms duration at 200 ms interval. We refer to this procedure as electrotransfer. The efficiency of the procedure was assessed by analyzing both the spread of GFP-linked fluorescence in the fin and by measuring its intensity. *Correspondence to: Sophie Vriz, Attache´ pour la science et la technologie, Ambassade de France, 4-11-44, Minami-Azabu, Minato-ku, Tokyo 1068514, Japan. E-mail:
[email protected] Grant sponsor: Le Ministe`re de la Recherche et l’Universite´ Paris 7, France.
28
TAWK ET AL.
FIG. 1. GFP expression in zebrafish caudal fin after DNA injection with (b–f) or without (a) the application of electric pulses. Fin specimens were observed with a fluorescence microscope. Ten g of pCMV-GFP were injected into the fin without electric administration (a) or submitted to electric pulses of 15 V (b). Fifteen g of DNA were injected into the fin, which was then submitted to eight electric pulses of 10 V (c). The fin received 15 g of DNA and submitted to 10 electric pulses of 15 V (d,e) and (f). Note that the signal is diffused extensively and strongly enhanced in fin tissues. Original magnification in (a– e) was 250⫻ and in (f) 400⫻.
FIG. 2. Fgf receptor 1 inhibition blocks fin regeneration and further outgrowth. (a– c): Fin injected with 15 g of pCMV-FGFR1 and submitted to 10 electric pulses of 15 V. The fin was cut 48 h after electrotransfer, and the regenerating part was examined 72 h postamputation. The fin showed no new growth. (b): Fin from untreated fish, 72 h after amputation, showing normal growth and new segmentation. (d): Fin injected with pCMV-GFP and submitted to electric pulses, 72 h postamputation showing normal morphology and regeneration. (e): Fin from fish allowed to regenerate for 72 h before treatment with pCMV-FGFR1 and analyzed 10 days after electrotransfer. Note that little outgrowth occurred during Fgf receptor 1 inhibition compared to untreated fin (f) where a complete regenerated fin is observed 10 days after amputation. Arrows demarcate amputation plane in each photograph. Original magnification in (a,b,d,f) was 7⫻, in (e) 10⫻ and in (c) 20⫻.
29
DNA ELECTROPORATION IN ADULT ZEBRAFISH FIN
Table 1 Effects of Voltage on the Efficiency of Gene Transfer and GFP Fluorescence* e m (nm)
exc ⫽ 490 nm Fluorescence intensity (a.u) 15 V 10 V 0V
500 505 510
35.1 11.4 5.0
11.0 3.6 1.6
8.8 2.8 1.3
15V/10V
Ratio 15V/0V
10V/0V
3.19 3.16 3.12
4.00 4.07 3.85
1.25 1.28 1.23
*Comparison of the fluorescence intensity of solutions issued from fins submitted to different electroporation parameters. All samples were diluted at the same protein concentration, which was determined using a Bradford essay. Fluorescence intensity values of each sample were corrected for autofluorescence by comparison with uninjected controls (internal standard control). ⫽ wavelength; exc ⫽ excitation; em ⫽ emission; a.u ⫽ arbitrary unit; V ⫽ volts.
Electrotransfer efficiency was found to depend on the characteristics of the electric pulses. The GFP expression was detected from a threshold field strength of 10 V, but too few cells were GFP positive at this voltage (Fig. 1c). In contrast, 20 V pulses caused extreme damage to fin tissues directly sandwiched between the electrode disks (data not shown). However, at 15 V, GFP expression was enhanced and a large number of cells showed a higher intensity in fluorescence signal (Fig. 1d, e, f). Varying number of pulses were tested, and the best results were obtained with 10 pulses (data not shown). The effect of injected DNA dosage on the efficiency of gene transfer was investigated by changing the amount of plasmid DNA injected. In the absence of electric pulses, no clear dose-effect relationship was observed because GFP fluorescence was generally weak. This was associated with rapid loss of the injected solution in the fins as a result of extensive swimming movements (Fig. 1a). We found that in the absence of electric pulses, GFP fluorescence was only detected at the original sites of injection. In contrast, the injected plasmid DNA diffused extensively in fin tissues after electroporation; cells positive for GFP were detected in the whole fin (Fig. 1f). When 10 g of DNA or less were injected and then electroporated, the number of GFP positive cells was small and signal was weak (Fig. 1b). An increase in the amount of injected DNA resulted in stronger signal intensity, it was much higher with 20 g than with 5 or 10 g (Fig. 1e). In order to provide a quantitative assessment of the gene transfer rate, we isolated total proteins from fin tissues of electroporated and nonelectroporated fins after pCMV-GFP injection (Table 1); a noninjected fin was used as a nonfluorescent negative control. We measured the fluorescence intensity of the GFP (excitation max.490 nm; ⬎ 90% efficiency at 488 nm), which is directly proportional to the number of fluorescent molecules in the extract. The results show that the electroporation at 15 V enhances the fluorescence signal four times in comparison to that of the injected nonelectroporated fins and three times to that of fins electroporated at 10 V (Table 1). We note a weak enhancement in the fluorescence signal in fins electroporated at 10 V in comparison to nonelectroporated fins, with a ratio of 1.3. Several measurements at different wavelengths (500, 505, 510) were compared in order to verify the
constancy of the established ratio (Table 1). All samples were diluted at the same protein concentration, determined using a Bradford assay. Fluorescence intensity values of each sample were corrected for autofluorescence by comparison with uninjected controls (Table 1). The time course of expression of electrotransferred pCMV-GFP was studied. GFP expression was detectable 48 h after DNA injection in the absence or presence of electric pulses and persisted for at least 15 days. Under these conditions, no alteration in fin morphology was observed after electrotransfer (Fig. 2d). Having established the optimal conditions to obtain high-efficiency DNA transfer into fin cells, we used this approach to disrupt Fgf signalling during fin regeneration. For this purpose, we used the pCMV-FGFR1 plasmid, which contains a dominant-negative form of the Xenopus Fgf receptor 1, lacking the intracellular tyrosine kinase domain. This truncated form of the receptor has been shown to strongly compromise Fgf signalling in Xenopus (Amaya et al., 1991). It has been previously shown that inhibition of this signalling pathway in the regenerating zebrafish caudal fin blocks blastema formation and its further outgrowth (Poss et al., 2000). We therefore tested the effect of the dominant-negative form of Fgf receptor 1 on blastema formation and blastema outgrowth in regenerating zebrafish caudal fin, by electrotransfer (10 pulses; 60 ms per pulse; 200 ms per interval; 15 V) of the construct either before or after amputation. Ten normal fins were injected and electroporated with 15 g of the pCMV-FGFR1; 48 h after electrotransfer, the fins were cut and animals were allowed to regenerate. Of the 10 fins examined 72 h postamputation, two regenerated normally, two showed regenerative defects, and six had a regenerative block (Fig. 2a– c). This suggests that the Fgf signalling pathway has been disrupted as expected. We also examined the effect of the pCMV-FGFR1 electrotransfer 72 h after fin amputation. Of the five fins examined 72 h postelectrotransfer, four showed a regenerative block for further outgrowth (Fig. 2e) and one had a normal regenerative outgrowth. These results further support the suggestion that the Fgf signalling pathway has been specifically targeted. These experiments appear to validate the electrotransfer approach to gene invalidation in the zebrafish.
30
TAWK ET AL.
Electrotransfer drastically increases the efficiency of gene transfer into fin tissues and is a suitable tool to obtain ectopic gene expression. Moreover, it gives longlasting expression and allows modulation of foreign gene expression by varying the amount of DNA injected, electric-pulse parameters, and/or the volume of tissues exposed to the electric pulses. These findings indicate that DNA electrotransfer in fin tissues may have broad applications for manipulating gene expression and analyzing the subsequent effects on cellular processes of fin regeneration. This will lead to a better understanding of the genetic mechanisms underlying epimorphic regeneration. METHODS Animals and Fin Amputations Zebrafish (Danio rerio) 3– 6 months of age were bought from SIDOLI (France). Wild-type fish were directly imported from India. Mature fish were kept at 28°C in photo-controlled aquariums and were anesthetized in neutralized methane sulfonate (MS 222, Sigma Chemical Co.) before cutting their fins. Animals were allowed to regenerate for various lengths of time; they were then anesthetized and the regenerated fin was removed for analyses. Plasmids and DNA Preparation The plasmids pCMV-GFP (pEGFP-C1, Clontech), and pCMV-FGFR1 (pCSXFD) (Amaya et al., 1991) contain the cytomegalovirus (CMV) promoter inserted upstream of the coding sequence, respectively, of the green fluorescent protein or of the Xenopus dominant negative form of fibroblast growth factor receptor l. Plasmids were prepared by using QIAGEN EndoFree Plasmid Maxi Kit as described by the manufacturer. DNA Injection and Electric-Pulse Delivery Different amounts of closed circular plasmid DNA, ranging from 5 to 15 g, were injected into the dermal skeleton of anesthetised adult zebrafish with microcapillaries. Ten fins were included in each experimental group. At defined times after DNA injection (25 s or 1 min, depending on the experiment), electric pulses were applied via a pair of electrode disks (7 mm diameter) rigged on the tips of tweezers (Suzuki et al., 1998) (pinsettes-Type electrode 520, Quantum BTX instrument). Pulses were delivered to each fin, which was placed between the electrode disks separated by a distance of approximately 2 mm. Electrical contact with the fin skeleton was ensured by applying a conductive gel (aquasonic 100 ultrasound transmission gel from Parker Laboratories, Inc.). Square-wave electric pulses were generated by an ECM 830 BTX electroporator (Genetronics, Inc.). Fish were then rapidly transferred to water at 28°C.
In Situ Detection of GFP Fins were removed at various times and mounted on glass sides with mounting medium (50% PBS pH 7.4, 50% glycerol). Green fluorescence was observed with a fluorescence microscope (Leitz, LABORLUX S) equipped with a WILD LEITZ camera (WILD MPS 52). Protein Extraction from Fins Total proteins were extracted from fins with the extraction buffer: 0.1 M Tris-Cl (pH 7.5), 0.2 M NaCl, 0,01 M  mercaptoethanol, 20% glycerol, and a protease inhibitor cocktail tablet used as described by the manufacturer (Roche Molecular Biochemicals). Chromosomal DNA was sheared by sonication. All debris was removed by centrifugation at 12,000 g for 5 min at 4°C, and the supernatant was transferred in a fresh tube. Three fins were used for each experiment. Optical Absorbance and Fluorescence Intensity Measurements Fluorescence measurements were recorded on an AMINCO BOWMAN spectrofluoremeter. The excitation and emission spectral bandwidths used were 4 nm and the optical pathlength was 1 cm. The excitation wavelength used was 490 nm, and the fluorescence intensity was recorded between 500 and 560 nm. All specimens were diluted similarly so that the optical density at 490 nm was less than 0.1. ACKNOWLEDGMENTS We thank Dr. Musci for providing pCMV-FGFR1 clone, Sandra Pellegrini and Dr. Jerome Collignon for comments on the manuscript, and Dr. Zentz for expert assistance in fluorescence measurement. LITERATURE CITED Aihara H, Miyazaki J. 1998. Gene transfer into muscle by electroporation in vivo. Nat Biotechnol 16:867– 870. Akamatsu W, Okano HJ, Osumi N, Inoue T, Nakamura S, Sakakibara S, Miura M, Matsuo N, Darnell RB, Okano H. 1999. Mammalian ELAV-like neuronal RNA-binding proteins HuB and HuC promote neuronal development in both the central and the peripheral nervous systems. Proc Natl Acad Sci U S A 96:9885–9890. Amaya E, Musci TJ, Kirschner MW. 1991. Expression of a dominant negative mutant of the FGF receptor disrupts mesoderm formation in Xenopus embryos. Cell 66:257–270. Ando H, Furuta T, Tsien RY, Okamoto H. Photo-mediated gene activation using caged RNA/DNA in zebrafish embryos. Nat Genet 28: 317–325. Brockes JP. 1997. Amphibian limb regeneration: rebuilding a complex structure. Science 276:81– 87. Eide FF, Eisenberg SR, Sanders TA. 2000. Electroporation-mediated gene transfer in free-swimming embryonic Xenopus laevis. FEBS Lett 486:29 –32. Geraudie J, Ferretti P. 1998. Gene expression during amphibian limb regeneration. Int Rev Cytol 180:1–50. Goto T, Nishi T, Tamura T, Dev SB, Takeshima H, Kochi M, Yoshizato K, Kuratsu J, Sakata T, Hofmann GA, Ushio Y. 2000. Highly efficient electro-gene therapy of solid tumor by using an expression plasmid for the herpes simplex virus thymidine kinase gene. Proc Natl Acad Sci U S A 97:354 –359.
DNA ELECTROPORATION IN ADULT ZEBRAFISH FIN
Johnson SL, Weston JA. 1995. Temperature-sensitive mutations that cause stage-specific defects in Zebrafish fin regeneration. Genetics 141:1583–1595. Mir LM, Bureau MF, Gehl J, Rangara R, Rouy D, Caillaud JM, Delaere P, Branellec D, Schwartz B, Scherman D. 1999. High-efficiency gene transfer into skeletal muscle mediated by electric pulses. Proc Natl Acad Sci U S A 96:4262– 4267. Muramatsu T, Mizutani Y, Ohmori Y, Okumura J. 1997. Comparison of three nonviral transfection methods for foreign gene expression in early chicken embryos in ovo. Biochem Biophys Res Commun 230:376 –380. Nasevicius A, Ekker SC. 2000. Effective targeted gene ’knockdown’ in zebrafish. Nat Genet 26:216 –220. Poss KD, Shen J, Nechiporuk A, McMahon G, Thisse B, Thisse C,
31
Keating MT. 2000. Roles for Fgf signaling during zebrafish fin regeneration. Dev Biol 222:347–358. Santamaria JA, Becerra J. 1991. Tail fin regeneration in teleosts: cellextracellular matrix interaction in blastemal differentiation. J Anat 176:9 –21. Suzuki T, Shin BC, Fujikura K, Matsuzaki T, Takata K. 1998. Direct gene transfer into rat liver cells by in vivo electroporation. FEBS Lett 425:436 – 440. Swartz M, Eberhart J, Mastick GS, Krull CE. 2001. Sparking new frontiers: using in vivo electroporation for genetic manipulations. Dev Biol 233:13–21. White JA, Boffa MB, Jones B, Petkovich M. 1994. A zebrafish retinoic acid receptor expressed in the regenerating caudal fin. Development 120:1861–1872.
