Increased level and duration of expression in muscle by co-expression of a transactivator using plasmid systems

Abstract

Skeletal muscle is an attractive target for gene therapies to treat either local or systemic disorders, as well as for genetic vaccination. An ideal expression system for skeletal muscle would be characterized by high level, extended duration of expression and muscle specificity. Viral promoters, such as the cytomegalovirus (CMV) promoter, produce high levels of transgene expression, which last for only a few days at high levels. Moreover, many promoters lack muscle tissue specificity. A muscle-specific skeletal α-actin promoter (SkA) has shown tissue specificity but lower peak activity than that of the CMV promoter in vivo. It has been reported in vitro that serum response factor (SRF) can stimulate the transcriptional activity of some muscle-specific promoters. In this study, we show that co- expression of SRF in vivo is able to up-regulate SkA promoter-driven expression about 10-fold and CMV/SkA chimeric promoter activity by five-fold in both mouse gastrocnemius and tibialis muscle. In addition, co-expression of transactivator with the CMV/SkA chimeric promoter in muscle has produced significantly enhanced duration of expression compared with that shown by the CMV promoter-driven expression system. A dominant negative mutant of SRF, SRFpm, abrogated the enhancement to SkA promoter activity, confirming the specificity of the response. Since all the known muscle-specific promoters contain SRF binding sites, this strategy for enhanced expression may apply to other muscle-specific promoters in vivo.

Introduction

Skeletal muscle is an attractive tissue for non-viral gene delivery for many reasons, which include the following. Large quantities of skeletal muscle are available and accessible in animals for intramuscular injection. Muscle fibers are capable of taking up naked plasmid DNA after intramuscular adminstration.1,2 Exogenous plasmids in muscle cells have produced expression of transgene proteins for more than 1 year at low expression level.1,2,3 Muscle can function as a ‘bioreactor’ to treat regional and systemic disease.4 Finally, formulated plasmids or electroporation can lead to a large percentage of muscle cells being transfected, high level expression and excellent reproducibility.5,6

Transgene expression in skeletal muscle has been demonstrated using a variety of plasmid backbones and reporter genes driven by a number of viral promoters. These promoters include the cytomegalovirus (CMV) immediate–early promoter,7,8,9 Rous sarcoma virus (RSV) long terminal repeat (LTR) promoter,1 and the SV40 early promoter.10 However, these viral promoters may be inactivated in muscle tissue.11 Moreover, not all of the administered plasmid remains in the muscle. An uncontrolled expression of a therapeutic gene in non-target cells could potentially lead to undesirable local or systemic side-effects.8 An avian skeletal muscle tissue-specific α-actin promoter (SkA) has been employed to produce tissue-specific expression.12 This promoter has been well-characterized and is known to be active in adult skeletal muscle tissues.12,13 Using this promoter, biologically active human insulin-like growth factor-I (hIGF-I) and human growth hormone (hGH) have been produced in murine muscle.8,9 One potential limitation of the SkA promoter to gene therapy applications is that, in general, it yields lower levels of expression in vivo than the stronger CMV viral promoter.

Barnhart et al14 combined a number of enhancers and promoters from various muscle-specific genes with the strong CMV gene enhancer/promoter to generate a more active promoter. Only one out of the 20 combinations showed better activity than that of the CMV promoter. Nettelbeck et al15 recently reported a strategy to enhance tissue-specific promoter activity in vitro by using the viral transactivator VP16. We have screened several non-viral muscle-specific promoter transactivators (Figure 1) to enhance SkA promoter transcriptional activity in muscle in vivo. In this article, we report that co-expression of a serum response factor (SRF) in murine muscle not only enhanced SkA promoter activity about 10-fold and a CMV/SkA chimeric promoter activity by five-fold, but also prolonged the duration of expression for more than 21 days.

Figure 1
figure1

Gene constructs. All constructs contain the Valentis backbone with the characteristic elements of a UT12 (5′ UTR sequence) and a downstream intron (IVS8) (Valentis, Inc, unpublished). Luciferase or human placental alkaline phosphatase (SEAP) cDNA was used as reporter genes. The pLC1072 plasmid contains a muscle-specific promoter, the skeletal muscle α-actin promoter (SkA). All constructs contain a human growth hormone poly(A) signal (3′ hGH), except for pLC0888, which contains a bovine growth hormone poly(A) signal (3′ bGH). pLC0888 and pAP1166 plasmids contain a viral CMV promoter. pLC1193 and pAP1396 plasmids contain a chimeric of the CMV enhancer, from −238 to −522 relative to the transcription start site, and an SkA promoter fragment, −119 to −499. pLC1072 and pLC0888 plasmids code for luciferase and pAP1166 and pAP1396 plasmid code for SEAP.

Results

Muscle-specific SkA promoter (pLC1072) is active in vitro but less in vivo

Two expression systems with the luciferase reporter gene, one driven by muscle-specific SkA promoter (pLC1072) and the other driven by non-muscle specific CMV promoter (pLC0888), were characterized in vitro and in vivo. In chicken primary myotubes in vitro, both the muscle-specific SkA and non-specific CMV-driven systems yielded high luciferase activity (Figure 2a), 1.7 × 1010 and 9.9 × 109 rlu/mg protein, respectively.

Figure 2
figure2

Comparison of transcriptional activity of viral promoter CMV and muscle promoter SkA in vitro (a) and in vivo (b) using luciferase. The activity of luciferase was measured 2 days after transfection in vitro or administration in vivo in murine tibialis muscles. Mean and standard error are shown in the Figure (n = 10 tibialis muscles). One microgram plasmid was used per well in vitro and 12.5 μg per tibialis muscle in vivo.

To obtain a rapid and strict comparison of the expression level for these two expression systems in vivo, samples were harvested 2 days after the plasmid injection to muscle. Differences in expression levels in vivo were observed. Lower luciferase activity was observed both in mouse tibialis cranialis and gastrocnemius muscle in vivo with the SkA-driven expression system (pLC1072), as compared with the strong viral CMV-driven system (pLC0888). Typically, the CMV-driven expression system (pLC0888) showed 200- to 500-fold higher peak luciferase activity than the SkA promoter expression system (pLC1072), 2 days after adminstration of plasmids in the tibialis muscle (Figure 2b). This result was reproducibly observed in three independent experiments.

Co-expression of SRF activates the SkA promoter in murine muscle tissue in vivo

It has been reported in vitro that SRF alone can give a two-fold enhancement of the transcriptional activity of the SkA promoter in differentiation-blocked myoblasts,16 as well as that of the cardiac α-actin promoter in a fibroblastic 10T1/2 cell.17 We have determined that co-expression of SRF with the SkA promoter stimulated the SkA promoter activity in murine muscle in vivo. The level of activation achieved by co-expression of SRF was about five- to 14-fold higher than that of SkA alone in both murine gastrocnemius and tibialis cranialis muscles on day 2 (P < 0.05) (Figure 3). In three independent experiments, the average increase with SRF co-expression was about 10-fold higher than SkA alone. These results indicate that a single transcription factor, SRF, can enhance muscle-specific promoter activity in different muscle tissues in vivo.

Figure 3
figure3

Effect of co-expression of serum response factor (SRF) on SkA promoter activity in murine gastronemius and tibialis muscle. Twenty micrograms of SkA-luciferase plasmid (pLC1072) and 5 μg of SRF expression plasmid (pSRF) or the control plasmid (pCGN) were co-formulated in 50 μl of saline and injected into gastrocnemius muscles. Half of the same plasmid dose was administered to the tibialis muscles of the same mice. Luciferase activity in the muscles was determined 2 days after injection. Each bar is the mean ± s.e.m (n = 10 muscles).

Up-regulation of SkA promoter activity by SRF is SRF-binding dependent

It is known that CArG elements, also known as muscle response elements (MRE) or serum response elements (SRE), are present in all muscle-specific promoters. These elements function as SRF binding sites and these SRF binding sites are sufficient for muscle-specific expression.18 To evaluate if SRF binding to the SkA promoter is essential for enhancement of this promoter activity, an SRF dominant-negative mutant form (SRFpm) expression plasmid was co-administered with SkA-driven luciferase expression construct (pLC1072) in gastrocnemius muscle. This mutant has three point mutations in the DNA binding domain of SRF that converts three critical base contact amino acids, Arg143, Lys145 and Leu146, to the neutral amino acids, ILU143, ALA145 and GLY146, respectively.19 These mutations abolish the DNA binding activity of the mutant form SRF, SRFpm. As expected, no enhancement of luciferase activity was observed when SkA-driven luciferase (pLC1072) was co-expressed with pSRFpm (Figure 4a). The loss of DNA binding activity of SRFpm totally abrogated augmentation of SkA activity. A truncated form of SRF (SRF1–266), in which the C-terminal activation domain was deleted and the DNA binding and dimerization domain were preserved, enhanced SkA promoter activity only moderately (Figure 4a). These results suggest that binding of functional SRF to the promoter is critical for promoter activation.

Figure 4
figure4

Mechanism of SRF-enhanced effect on SkA promoter activity. (a) Dependence of SRF binding for enhancement of SkA promoter activity. Twenty micrograms of SkA-luciferase (pLC1072) plasmid was co-formulated with 5 μg of other forms of SRF or muscle-specific transcription factor expression plasmids (MyoD or Myogenin) in 50 μl of saline and injected into gastrocnemius muscle (n = 10 muscles). Each bar represents the mean ± s.e.m. of 10 muscles. The different alphabetical letters denote statistically significant differences (P < 0.05). (b) SRF dose-dependent enhancement on SkA promoter activity. Ten micrograms of SkA-luciferase plasmid (pLC1072) mixed with varying doses of the SRF expression plasmid was formulated in 25 μl of saline, and injected into murine tibialis muscle (n = 10 muscles). The control plasmid (pCGN) was used to maintain a constant mass of DNA. Each bar represents the mean ± s.e.m.

To test further the hypothesis that SRF binding to SkA promoter is essential for enhancement of the promoter activity, we varied the ratio of SRF expression plasmid (pSRF) to the SkA-driven luciferase plasmid (pLC1072). A 10 μg dose of SkA-luciferase (pLC1072) was formulated with various amounts of SRF expression plasmid (pSRF) ranging from 0.5 to 5 μg. To keep the total DNA amounts administered constant (15 μg) for each murine tibialis muscle, an ‘empty plasmid’ with no coding sequence (pCGN) was included in the formulation as needed. Luciferase activity of the murine tibialis muscle tissue was determined 2 days after injection (Figure 4b). As the amount of pSRF decreased, luciferase expression driven by SkA promoter also decreased. The weight ratio of SRF expression plasmid to SkA-driven expression plasmid for optimal activation was 1:4. The dose dependence of SRF for the activation of SkA promoter further supports the hypothesis that this is a truly functional SRF effect.

Duration of SRF trans-activation of SkA promoter

We examined the duration of expression by activating the SkA promoter with SRF after a single dose administration. After administration of 20 μg of SkA-luciferase (pLC1072) formulated with 5 μg of pSRF (or pCGN for the control) plasmid, luciferase activity was assayed at 2, 7, 14 and 28 days (Figure 5). The relative up-regulation of SkA activity by SRF co-expression continued for 28 days. The maximum up-regulation was observed on day 7 and declined thereafter (Figure 5a). These data indicate that such a marked up-regulation was a persistent effect and that the intensity of up-regulation was time-dependent. The SRF effect declined with time after injection. This decline corresponds to the decline of SkA promoter activity in muscle.

Figure 5
figure5

Effect of co-expression of SRF on duration of gene expression under control of the SkA promoter. Twenty micrograms of SkA-luciferase (pLC1072) and 5 μg SRF expression plasmid or control plasmid (pCGN) were formulated in 50 μl saline and injected into murine gastrocnemius muscles (n = 10 muscles). Each bar represents mean ± s.e.m. (a) SkA-luciferase with co-expression of SRF (pLC1072+ pSRF, –♦–) showed better duration of expression than did SkA-luciferase alone (pLC1072+pCGN, –▪–). (b) CMV-luciferase (pLC0888, –•–) was used as a control.

Co-expression of SRF with a CMV/SKA chimeric promoter-driven system in muscle exceeds the expression level and duration of expression of CMV-driven expression system

Although SkA promoter activity was enhanced by co-expression of SRF, the gene expression level driven by this promoter was still lower than that produced by CMV promoter-driven system (Figure 4). To develop an expression system with CMV promoter-driven expression level but with longer duration, a CMV/SkA chimeric promoter-luciferase (pLC1193) and -SEAP were constructed (pAP1396, Figure 1). We then examined whether this promoter could increase the expression level in muscle and retain the up-regulation responsiveness to SRF in vivo. Both luciferase (pLC1193) and SEAP (pAP1396) reporter genes were used in this experiment. SEAP was chosen as the reporter gene to permit a time-course study by measurement of expressed protein secreted in serum with each animal. Luciferase (pLC1193) was measured at day 2 for comparison with the previous experiment. SRF co-expression enhanced CMV/SkA chimeric promoter activity to 40% of that produced by CMV promoter on day 2 (Figure 6a). However, the expression level of SEAP driven by CMV/SkA chimeric promoter with co-expression of SRF was as high as CMV expression system on day 7, and greater than that of CMV on days 14 and 21 (Figure 6b). Co-expression of SRF with a chimeric promoter not only increased expression levels, but also maintained expression levels up for at least 21 days.

Figure 6
figure6

Effect of co-expression of SRF on chimeric and CMV promoter activity. Twenty micrograms of either CMV/SkA-luciferase (pLC1193), CMV/SkA-SEAP (pAP1396) or CMV-luciferase (pLC0888) plasmid and 5 μg of SRF expression plasmid or control plasmid were formulated in 50 μl of saline and injected into murine gastrocnemius muscles (n = 10 muscles). Each bar represents mean ± s.e.m. (a) Effect of co-expression of SRF on the level of gene expression under the control of chimeric promoter. Black and white bars represent with and without co-expression of SRF, respectively. The different alphabetical letters indicate statistically significant differences. CMV promoter-driven luciferase gene (pLC0888) was used as a control. (b) Effect on duration of gene expression under control of chimeric promoter with co-expression SRF. CMV-SEAP expression system (pAP1166 –♦–) showed highest expression at day 3 after administration of plasmid. CMV/SkA-SEAP with co-expression of SRF (pAP1396+pSRF –▪–) showed better expression duration and level than either CMV/SkA-SEAP alone (pAP1396+pCGN, ––) or the CMV-SEAP system. (c) Evidence for promoter-specific SRF effect. Twenty micrograms of CMV-luciferase (pLC0888) mixed with 5 μg of pSRF (or control plasmid pCGN) in 50 μl of saline were injected into CD1 murine gastrocnemius muscles. Ten muscles were used for each treatment. Each bar represents the mean ± s.e.m. of 10 muscles. No statistically significant differences were found.

CMV promoter without SRF binding sites did not respond to SRF co-expression in vivo

It has been well documented that SRF binds to the serum response element (SRE) in the promoter region and thereby regulates gene expression.16 As expected, expression levels with a CMV promoter-driven system, which does not have any SRF binding site, were not changed by co-expression with SRF (Figure 6c). These results suggest that SRF effect is promoter context- specific, with a requirement for a SRE in the promoter.

Discussion

It has been clearly demonstrated in previous studies that a skeletal muscle-specific SkA promoter exhibits strong specificity of gene expression in muscle in vivo.13 However, the expression levels of SkA promoter-driven system were much lower than that of the CMV promoter-driven expression system in vivo (Figure 2b). This observation is also true for a number of other tissue-specific promoters, such as the endothelium-specific promoter.15 To enhance the SkA promoter activity, we used two strategies. The first is the addition of a cis-element upstream of this promoter and the second is by co-expression of a trans-activator.

We have shown that CMV/SkA chimeric promoter (pLC1093 and pAP1396) produced 10% activity of the CMV promoter-driven system in vivo 2 days after injection. When an SRF trans-activator was co-expressed in vivo, this chimeric promoter activity was stimulated further (Figure 6a). Even on day 2–3, when the expression driven by the SkA promoter was relatively low, expression driven by the CMV/SkA chimeric promoter along with co-expression of SRF was about 40–70% of the CMV promoter-driven expression (Figure 6). This combination also produced higher expression levels than those of the CMV-SEAP system at day 14 and 21. More importantly, co-expression of SRF with either SkA (pLC1072) or CMC/SkA chimeric promoter (pLC1093, pAP1396) produced longer duration of expression. Thus, a combination of cis-element improvements, as well as the application of a positive transactivator such as SRF, offers a means for increasing both levels and duration of gene expression.

SRF is known as a ubiquitous and conserved transcription factor. It is a key regulator of cellular response genes termed immediate–early genes, such as Egr-1 and Egr-2,20 and non-muscle actin.21 The cis elements with a CArG (CC(A/T)6GG) sequence motif mediate a rapid and transient serum inducibility of these genes. More importantly, SRF has been found to be critical in muscle-specific gene expression and muscle development.16,17,18,19,22,23,24,25,26 The SRF binding motif is found within the promoters of many, if not all, important muscle-specific genes, including actin, myosin heavy chains, myosin light chain, muscle creatine kinase, dystrophin, troponin T and SRF genes. YY1, a zinc finger transcription factor,27 also binds to the skeletal muscle α-actin promoter SRE1,16,23 and competes with SRF for binding to these SREs. Overexpression of YY1 represses these elements, and SRF overexpression is able to reverse this repression.28 Accordingly, expression of SRF in vivo would be expected to compete with YY1 and perhaps other negative trans-regulators, such as F-ACT116 and thus in favor of more active transcription. We do not have direct nuclear run-on data to support this hypothesis, because no technique is yet available for isolating nuclei from muscle tissue. Quantitative RT-PCR data did show a five-fold increase in the copy number of steady-state insulin-like growth factor-I (IGF-I, pIG0552 driven by SkA promoter, Alila et al8) mRNA driven by SkA promoter with co-expression of SRF in murine muscle (data not shown). This suggests that the likely mechanism of SRF enhancement on SkA- and CMV/SkA promoter-driven gene expression is to enhance the promoter transcriptional activity.

Materials and methods

Plasmid construction and preparation

Six constructs with four different promoters and two reporter genes were constructed in the Valentis’s plasmid backbone, which includes 107 bp of 5′UTR (UT12), a 117 bp of 5′ synthetic intron (ivs8), a kanamycin resistance gene and a PUC12 backbone. Plasmid pLC1072, pLC0888 and pLC1193 all contained a luciferase reporter gene but different promoters (Figure 1). Constructs pAP1396 and pAP1166 had the same backbone as pLC1193 and pLC0888, but a human placental alkaline phosphatase reporter gene was used. The promoters used in various constructs were an SkA promoter,9,29 a CMV enhancer/promoter and a CMV/SkA chimeric promoter. The SkA promoter (pLC1072) was obtained by performing PCR on pIG05529 plasmid using sense primer 5′-AGCCTGCAGGGGCCGCTCTAGCTAGAGTCT-3′ and antisense primer 5′-AGCTCGAGCCTGTGCTGACTGCG CGTCG-3′. The 518-bp PCR product was cut with Sse8387 and XhoI, introduced in the primer and subcloned into the Valentis’s backbone cut with the same restriction enzymes. For pLC0888, the CMV enhancer/promoter was excised from pCMV-hGH-GH7 and subcloned into standard Valentis’s plasmid. The CMV/SkA chimeric promoter construct (pLC1193) was made by subcloning the CMV enhancer element cut with Sse8387I and SnaBI into pLC1072 cut with Sse8387I and StuI. SRF, SRF mutant and truncated SRF expression plasmids, pSRF, pSRFpm and pSRF1–266, respectively, were described in a previous publication.19 The control plasmid pCGN was made by the deletion of SRF cDNA from pSRF. All SRF-related plasmids contained a CMV promoter, HSVtk 5′ leader, an HA epitope and a 3′ rabbit β-globin intron and polyadenylation site.

All plasmids were manufactured using Qiagen Giga Endotoxic Free Prep Kit (Valencia, CA, USA). Residual salts were removed from the plasmid by dialysis against sterile water (USP, Baxter Deerfield, IL, USA) with a Millipore (Bedford, MA, USA) dialysis tube. The purity of the plasmid was confirmed by 1% agarose gel electrophoresis. DNA concentration was measured by absorption at 260 nm. The percentage of supercoiled DNA to the total DNA and the OD 260/280 ratios of these plasmid preparations were in the range of 80–90% and 1.7–1.9, respectively.

Formulation and intramuscular injection of DNA plasmid

Formulations were made by mixing plasmid, sterile water and 5 M NaCl to a final concentration of 150 mM. The formulation osmolarity was measured using a Fiske One-Ten Micro-Sample Osmometer (Fiske, Norwook, MA, USA).

Animals were anesthetized by intraperitoneal administration of a mixture of ketamine (42.8 mg/ml), xylazine (8.6 mg/ml) and acepromazine (1.4 mg/ml) at a dose of 0.7–1.0 ml/kg. The tibialis and gastrocnemius muscles of anesthetized 6–8-week-old CD-1 mice (Charles River, Wilmington, MA, USA) were injected intramuscularly with 25 and 50 μl of formulation containing 12.5 and 25 μg DNA, respectively, using a 28½ G needle. The mice were kept at 37°C until they regained consciousness. The animals were killed by CO2 asphyxiation. Tibialis cranialis and gastrocnemius muscles were harvested, transferred into an Eppendorf tube, immediately frozen in liquid nitrogen and lyophilized overnight.

Protein extraction and luciferase activity assay

Frozen lyophilized muscles were homogenized using mini bead-beater (Biospec Products, Bartlesville, OK, USA) with silica beads for 2 min. One milliliter of luciferase cell lysis buffer (Promega, Madison, WI, USA) was added to the powdered muscle and the samples were homogenized for another 3 min. The suspension was centrifuged at 1500 g for 15 min, and the supernatant was used to assay total protein and luciferase activity. Luciferase activity was assayed using a microplate luminometer (Wallac, Gaithersburg, MD, USA). One hundred microliters of luciferase substrate was automatically added to 20 μl of muscle extract (previously diluted 1:2) and relative light units were recorded. The total protein was determined with the BCA protein assay kit (Pierce, Rockford, IL, USA).

Statistical analysis

Experimental data were analyzed by one-way analysis of variance with measurement of relative light units as the main effect. Means of individual treatment were compared using a Student’s t test when the main effect was significant. Statistical significance was defined as a P < 0.05.

References

  1. 1

    Wolff JA et al. Direct gene transfer into mouse muscle in vivo Science 1990 247: 1465–1468

  2. 2

    Wolff JA et al. Long-term duration of plasmid DNA and foreign gene expression in mouse muscle Hum Mol Genet 1992 1: 363–369

  3. 3

    Danko I et al. Pharmacological enhancement of in vivo foreign gene expression in muscle Gene Therapy 1994 1: 114–121

  4. 4

    Coney L et al. Facilitated DNA inoculation induces anti-HIV-1 immunity in vivo Vaccine 1994 12: 1545–1550

  5. 5

    Aihara H, Miyazaki JI . Gene transfer into muscle by electroporation in vivo Nature Biotechnol 1998 16: 867–870

  6. 6

    Mumper RJ et al. Protective interactive noncondensing (PINC) polymers for enhanced plasmid distribution and expression in rat skeletal muscle J Control Rel 1998 52: 191–203

  7. 7

    Tripathy SK et al. Long-term expression of erythropoietin in the systemic circulation of mice after intramuscular injection of a plasmid DNA vector Proc Natl Acad Sci USA 1996 93: 10876–10880

  8. 8

    Alila H et al. Expression of biologically active human insulin-like growth factor-I following intramuscular injection of a formulated plasmid in rats Hum Gene Ther 1997 8: 1785–1795

  9. 9

    Anwer K et al. Systemic effect of human growth hormone after intramuscular injection of a single dose of a muscle-specific gene medicine Hum Gene Ther 1998 9: 659–670

  10. 10

    Wells DJ, Goldspink G . Age and sex influence expression of plasmid DNA directly injected into mouse skeletal muscle FEBS Lett 1992 306: 203–205

  11. 11

    Dai Y, Roman M, Naviaux RK, Verma IM . Gene therapy via primary myoblasts: long-term expression of factor IX protein following transplantation in vivo Proc Natl Acad Sci USA 1992 89: 10892–10895

  12. 12

    Chow KL, Schwartz RJ . A combination of closely associated positive and negative cis-acting promoter elements regulates transcription of the skeletal α-actin gene Mol Cell Biol 1990 10: 528–538

  13. 13

    Petropoulos CP et al. The chicken skeletal muscle α-actin promoter is tissue specific in transgenic mice Mol Cell Biol 1989 9: 3785–3792

  14. 14

    Barnhart KM et al. Enhancer and promoter chimeras in plasmids designed for intramuscular injection: a comparative in vivo and in vitro study Hum Gene Ther 1998 9: 2545–2553

  15. 15

    Nettelbeck DM, Jerome V, Muller R . A strategy for enhancing the transcriptional activity of weak cell type-specific promoters Gene Therapy 1998 5: 1656–1664

  16. 16

    Lee TC, Chow KL, Fang P, Schwartz RJ . Activation of skeletal α-actin gene transcription: the cooperative formation of serum response factor-binding complexes over positive cis-acting promoter serum response elements displaces a negative-acting nuclear factor enriched in replicating myoblasts and nonmyogenic cells Mol Cell Biol 1991 11: 5090–5100

  17. 17

    Chen CY, Schwartz RJ . Competition between negative acting YY1 versus positive acting serum response factor and tinman homologue Nkx-2.5 regulates cardiac α-actin promoter activity Mol Endocrinol 1997 11: 812–821

  18. 18

    Walsh K . Cross-binding of factors to functionally different promoter elements in the c-fos and skeletal actin genes Mol Cell Biol 1989 9: 2191–2201

  19. 19

    Croissant JD et al. Avian serum response factor expression restricted primarily to muscle cell lineages is required for α-actin gene transcription Dev Biol 1996 177: 250–264

  20. 20

    Gius D et al. Transcriptional activation and repression by Fos are independent functions: the C terminus represses immediate–early gene expression via CarG elements Mol Cell Biol 1990 10: 4243–4255

  21. 21

    Miwa T et al. Structure, chromosome location and expression of the human smooth muscle (enteric type) gamma-actin gene: evolution of six human actin genes Mol Cell Biol 1991 11: 3296–3306

  22. 22

    Gilman MZ, Wilson RN, Weinerg RA . Multiple protein-binding sites in the 5′-flanking region regulate c-fos expression Mol Cell Biol 1986 6: 4305–4316

  23. 23

    Lee TC, Shi Y, Schwartz RJ . Displacement of BrdUrd-induced YY1 by serum response factor activates skeletal α-actin transcription in embryonic myoblasts Proc Natl Acad Sci USA 1992 89: 9814–9818

  24. 24

    Chen CY et al. Activation of the cardiac α-actin promoter depends upon serum response elements Dev Genet 1996 19: 119–130

  25. 25

    Gauthier-Rouviere C et al. Expression and activity of serum response factor are required for muscle determining factor MyoD in both dividing and differentiating mouse C2C12 myoblasts Mol Biol Cell 1996 7: 719–727

  26. 26

    Belaguli N, Schildmeyer L, Schwartz RJ . Organization and myogenic restricted expression of the murine serum response factor gene J Biol Chem 1997 272: 18222–18231

  27. 27

    Martin KA et al. A competitive mechanism of CarG element regulation by YY1 and SRF: implications for assessment of Phox1/Mhox transcription factor interactions at CarG elements DNA Cell Biol 1997 16: 653–661

  28. 28

    Gualberto A et al. Functional antagonism between YY1 and the serum response factor Mol Cell Biol 1992 12: 4209–4212

  29. 29

    Coleman ME et al. Myogenic vector expression of insulin-like growth factor I stimulates muscle cell differentiation and myofiber hypertrophy in transgenic mice J Biol Chem 1995 270: 12109–12116

Download references

Acknowledgements

We thank Dr Ron Brywes’s laboratory for some of SRF constructs and Alain Rolland, Mike Fons and Sean Sullivan for their critical reading of the manuscript.

Author information

Correspondence to L C Smith.

Rights and permissions

Reprints and Permissions

About this article

Cite this article

Li, S., MacLaughlin, F., Fewell, J. et al. Increased level and duration of expression in muscle by co-expression of a transactivator using plasmid systems. Gene Ther 6, 2005–2011 (1999). https://doi.org/10.1038/sj.gt.3301032

Download citation

Keywords

  • gene expression
  • trans-activation
  • SRF
  • duration
  • muscle
  • plasmid

Further reading