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. Author manuscript; available in PMC: 2021 Oct 14.
Published in final edited form as: Syst Biol Reprod Med. 2018 Aug 23;64(5):389–398. doi: 10.1080/19396368.2018.1499153

Antisense techniques provide robust decrease in GnRH receptor expression with minimal cytotoxicity in GT1-7 cells

Maurice Andre Recanati a, Hongling Du a, Katherine J Kramer b, Maik Hüttemann c, Robert A Welch a
PMCID: PMC8515877  NIHMSID: NIHMS1595386  PMID: 30136857

Abstract

The episodic pattern of gonadotropin-releasing hormone (GnRH) secretion from the hypothalamus is driven by an integrated network of cells termed the GnRH pulse generator. Cultured and immortalized GnRH neurons also produce a pulsatile pattern of GnRH secretions when grown in the absence of other cell types, suggesting the presence of an intrinsic oscillator mediating GnRH secretion. The mechanisms underlying such pulsatility comprise one of the most tantalizing problems in contemporary neuroendocrinology. In order to study the mechanism by which GnRH is produced in a pulsatile fashion, the autocrine effect of GnRH on GnRH-producing neurons must be eliminated. This may be performed by downregulating the expression of the GnRH receptor. Treatment with three 21-mer exogenous phosphorothioates and transient transfections with an inducible plasmid containing an antisense construct to the GnRH receptor gene decreased GnRH receptor expression further. This resulted in less cytotoxicity compared to inhibition of RNA or protein synthesis with actinomycin D, α-amanitin, puromycin, and cycloheximide. This study shows methods and optimized conditions established for the generation of a stable GT1–7 cell line containing an inducible construct allowing the downregulation of GnRH receptor expression.

Keywords: GnRH, GnRH pulse generator, antisense inhibitors, GT1-7 cells, inducible plasmid

Introduction

Reproduction in mammals is regulated by the interplay of three primary endocrine organs: the hypothalamus, the anterior pituitary, and the gonads. The major humoral regulator of this reproductive axis, gonadotropin-releasing hormone (GnRH), is produced by neuronal cells located in the preoptic area and adjacent sites in the rostral portion of the hypothalamus and secreted into the hypophyseal portal vessels at the median eminence (Silverman 1994; Levy et al. 2005).

The ‘hypothalamic pulse generator’ discovered by Ernst Knobil consists of approximately 1,500 neurons in the arcuate nucleus of the thalamus in rhesus monkeys (Knobil 1990). However, location and cell numbers are species dependent (Wray and Hoffman 1986) (Wray 2010) (Rogers et al. 1998). Many clinical diseases (such as hypogonadotropic hypogonadism and Kallmann syndrome), physiological conditions (such as the onset of puberty), and contraceptives and infertility treatments (GnRH agonists and antagonists) are related to disruptions in the function of these specialized cells. In healthy individuals, the GnRH pulse generator that is ‘on’ in infancy, turns ‘off’ during childhood before switching back ‘on’ at puberty (Fritz and Speroff 2011). The mechanisms triggering the onset of puberty (Fritz and Speroff 2011), leading to associated disorders such as central precocious puberty, and activation of the hypothalamic-pituitary-gonadal axis (Anderson et al. 2003; Chumlea et al. 2003), delayed puberty from hypogonadotropic hypogonadism (Herman-Giddens et al. 1997) and Kallmann syndrome from failure of GnRH neuron migration during embryogenesis (Van Goor et al. 2000) remain poorly understood. Clinically, both GnRH agonists (leuprorelin) and GnRH antagonists (cetrotide) have been used to treat anovulatory infertility (Moghissi 2000) and increase the success of in vitro fertilization while minimizing the risks of ovarian hyperstimulation syndrome (Engmann et al. 2008; Humaidan and Alsbjerg 2014).

The secretion of the GnRH decapeptide in vivo takes place in a periodic fashion due to the activity of a hypothalamic GnRH pulse generator (Knobil 1981). Direct measurements on hypophyseal portal blood have demonstrated that GnRH release occurs in discrete bursts (Moenter et al. 1993), and that changes in the endocrine milieu such as endothelins, dopamine, adrenergic tone, glutamate, prolactin, GABA (Herbison and Moenter 2011; Constantin et al. 2013), estrogen (Hu et al. 2008; Chason et al. 2015), leptin (Woller et al. 2001), and even GnRH itself (Woller et al. 2004)) can modulate the frequency and amplitude of the secretory pulses (Drouva et al. 1983; Bedran de Castro et al. 1985; Krsmanovic et al. 1991; Roth et al. 2001). Furthermore, kisspeptin neurons within the hypothalamus directly contact GnRH neurons and can stimulate the release of GnRH through activation of the Kiss1 receptor within GnRH neurons (Sukhbaatar et al. 2013; Clarke et al. 2015; Maggi et al. 2016; Kanasaki et al. 2017b). In addition, GnRH metabolites, such as the pentapeptide GnRH-(1–5), have been found to regulate GnRH secretion through short loop feedback regulation. (Wu et al. 2005; Larco et al. 2015). Numerous in vitro observations have shown that primary cultures of hypothalamic neurons also secrete GnRH in a pulsatile manner, leading to the consensus that pulsatility is an inherent characteristic of these cells (Bourguignon and Franchimont 1981; Krsmanovic et al. 1992; Wetsel 1995). The precise mechanisms underlying how the pulsatile secretion and expression of GnRH itself are regulated are still largely unknown (Nunez et al. 1998; Kanasaki et al. 2017a). Clinically, in some forms of idiopathic hypogonadotropic hypogonadism, the GnRH neurons have developed and migrated properly but the GnRH pulse generating machinery is defective. A construct allowing GnRH-producing neurons to be easily grown in vitro, in the absence of external neural, endocrine and autocrine inputs, would facilitate studying the internal mechanisms responsible for pulsatile GnRH production.

The development of immortalized GnRH cells capable of secretory activity has provided a valuable model to study the mechanisms of GnRH secretion. The GT1–7 cell line was obtained from transgenic mice expressing a hybrid gene composed of the 5′-flanking region of the rat GnRH gene linked to the cDNA encoding the SV40 T-antigen (Mellon et al. 1990; Wetsel et al. 1992). These cells, which express neuronal markers, have a vast endoplasmic reticulum and Golgi apparatus, as well as secretory granules (Weiner et al. 1992) that permit them to secrete GnRH and its associated GAP peptide (Liposits et al. 1991) during perfusion and in static culture (Krsmanovic et al. 1992). The availability of this homogenous population of GnRH-producing cells facilitates the distinction between secretion controlled by endogenous channels and receptors and the modulation mediated by connecting neurons. Comparative studies on GT1–7 cells and fetal hypothalamic cultures in vivo and in situ have demonstrated that the activity of the transformed cell line is controlled by multiple factors in a way that is comparable to primary hypothalamic cells (Stojilkovic et al. 1994).

It has been shown that GnRH acts directly on immortalized GnRH neuronal cells to regulate its own release via a specific seven-transmembrane Gqproteincoupled receptor (Larco et al. 2015). The cellular mechanism underlying this ultra-short feedback loop, however, remains unknown. In order to study the mechanisms responsible for the intrinsic GnRH pulse generation, it is necessary to eliminate the autocrine effect by reducing the expression of the GnRH receptor (GnRHR). Our first approach utilized pharmacological inhibitors (actinomycin D, α-amanitin, puromycin, and cyclophosphamide) to nonspecifically reduce expression of the GnRHR while subsequent approaches employed antisense constructs targeting expression of the receptor specifically. We evaluated the effect of these methods in reducing expression of the GnRHR and examined their cytotoxicity.

Results

Protein synthesis inhibitors

GT1–7 cells have been shown by binding studies and by RT-PCR to express the GnRHR that is capable of binding des-Gly10[D-Ala6]GnRH N-ethylamide, a radioiodinated competitive GnRH agonist, with a relatively high affinity (KD of about 0.9 nM versus 1.4 nM for the endogenous peptide) (Krsmanovic et al. 1999).

The biosynthetic inhibitors actinomycin D, cycloheximide, and puromycin were used to prevent cells from synthesizing new proteins and, because of receptor internalization, reduce the steady-state level of GnRHR expression (Figure 1). Using binding studies to quantify levels of the GnRHR, we found a 45% decrease in cell binding after a 24 h treatment with either cycloheximide or puromycin (p < 0.05), while actinomycin D caused only a 15% reduction. The decrease in binding was dose- and time-dependent since the 48 h treatment further reduced expression. A two-day incubation with puromycin reduced binding by nearly 65% (p < 0.05), while cycloheximide reduced it by about 55%.

Figure 1.

Figure 1.

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Effect of antineoplastic agents on GnRH receptor expression. Cells were incubated for 24 h in regular medium prior to the addition of antineoplastic agents at the following concentrations: 1 μg/mL actinomycin D, 10 μg/mL cycloheximide, and 100 μg/mL puromycin. Cells treated with cycloheximide or puromycin showed a statistically significant (*p < 0.05) decrease in GnRH receptor binding. Data from binding studies was adjusted to total cellular protein content and normalized to 24 h controls. Standard deviations are shown. Data represents four experiments (n = 4).

Amanitin treatment was shown to decrease binding by up to 40% in a dose-dependent fashion (Figure 2), but also caused cell detachment and cell death early in the experiment. It was found that a concentration of 50 μg/mL, delivered every other day, maximally inhibited receptor expression with optimal cell viability (p < 0.05).

Figure 2.

Figure 2.

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Effect of amanitin on receptor expression. Amanitin treatment, which causes a nonselective decrease in protein synthesis, decreased binding by about 40% at a concentration of 50 μg/mL (*p < 0.05). Data from binding studies was adjusted to total protein and normalized to controls (n = 4).

GnRHR knockdown using antisense oligonucleotides

The GnRHR targeted phosphorothioate oligonucleotides AS-1, AS-2, and AS-3 were used to treat GT1–7 cells. Specific binding of GnRH, an index of the quantity of GnRHR, was compared to the untreated controls and to cells treated with the randomly picked sense oligonucleotide as a control (Figure 3). The results showed a decrease in groups treated with antisense oligonucleotides. The oligonucleotide designated AS-2 showed a 60% decrease in binding (p < 0.05) and was more effective in downregulating binding than AS-1 (25%) or AS-3 (10%). Furthermore, the combination of AS-1 and AS-2, each at half the concentration of the single oligonucleotide, proved to act additively and decreased binding by up to 75% (p < 0.05). Five-day treatment with AS-2 alone or in combination with AS-1 significantly decreased specific binding when compared to the 48 h treatment (p < 0.05). In all three trials, groups treated with sense oligonucleotides exhibited a slightly lower level of binding relative to controls and appeared, on microscopic examination, to show some cell injury.

Figure 3.

Figure 3.

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Selective downregulation of the GnRH receptor using specific oligonucleotides. Three phosphorothioate oligonucleotides AS-1, AS-2, and AS-3 were used to treat GT1–7 cells and compared for their specific binding of GnRH (a surrogate of the quantity of GnRH receptor). Control cells were treated with a random sense oligonucleotide. AS-2 showed a 60% decrease in binding (*p < 0.05), and the combination of AS-1 and AS-2 proved to act additively and decreased binding by up to 75% (p < 0.05). Five-day treatment with AS-2 alone, or in combination with AS-1, significantly decreased specific binding when compared to 48 h treatment (p < 0.1 and < 0.05, respectively). Data from binding studies was adjusted to total protein and normalized to 48 h controls (n = 3).

Transient transfections

Transient transfections using the full-length antisense GnRHR transcript cloned into the Topo vector with the RXR plasmid showed a greater than 60% decrease in receptor expression (p < 0.05) when incubated in the presence of the cognate ligand, ponasterone A, compared to only a 10% drop in binding in its absence (Figure 4) when compared to non-transfected controls. There was no statistical difference between the 48 h treatment and the 5-day treatment groups. Transient transfections using ‘sense’ insert showed no difference when activated with ponasterone A, relative to non-transfected controls.

Figure 4.

Figure 4.

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Effect of transient transfection with RXR and full-length insert in the presence and absence of ecdysone. Transient transfections using the full-length antisense GnRH receptor transcript cloned into the Topo vector with the RXR plasmid showed a greater than 70% decrease in receptor expression (*p < 0.05) when incubated in the presence of the cognate ligand, ponasterone A (an insect steroid), compared to only a 15% drop in binding in its absence when compared to controls. This construct was not significantly more effective than the combination of AS-1 and AS-2 shown in Figure 3. There were no statistically significant differences at 48 h versus 5 days of treatment. Data from binding studies was adjusted to total protein and normalized to controls (n = 3).

Discussion

Protein synthesis inhibitors

GT1–7 cells were treated with antineoplastic agents to prevent cells from synthesizing new proteins. Actinomycin D inhibits transcription by binding DNA at the transcription initiation complex, preventing elongation of the nascent RNA chain by RNA polymerase. Cycloheximide acts by forming irreversible DNA interstrand and intrastrand crosslinks and puromycin inhibits translation by acting on the ribosome. Amanitin, a toxin extracted from the mushroom amanita phalloides, is a selective inhibitor of RNA polymerase II (Kedinger et al. 1970; Austoker et al. 1974). Because these antineoplastic agents act nonselectively to decrease overall protein synthesis and due to receptor internalization, a reduction in the steady-state level of GnRHR expression was observed. However, these compounds are toxic to dividing cells, such as GT1–7 cells, and caused a significant degree of detachment and morphological abnormalities. In addition, changes in cellular metabolic activity were suggested by a lack of acidification of the culture medium relative to the control group. The concentrations used in this paper represent maximal inhibition in receptor expression with optimal cell viability for this cell line.

Because of the high rate of cell death secondary to a nonspecific downregulation of protein synthesis, this pharmacological approach was extended in favor of a genetic approach capable of specifically targeting the GnRHR and having less cytotoxicity.

Antisense oligonucleotides

The decrease in binding shown with treatment by the three oligonucleotides is probably due to a decrease in receptor synthesis coupled with some level of receptor internalization. While the AS-2 oligonucleotide alone or in combination proved to be most effective, it is not known if this level of downregulation is sufficient to abolish the activity of the receptor on the GnRH secretion pattern and further testing is necessary.

In all three trials, groups treated with sense oligonucleotides exhibited a slightly lower level of binding relative to controls and appeared, on microscopic examination, to show some cell injury. This necrosis, which was more significant in cells treated with two doses of DNA over a 5-day course, is probably secondary to the dose-dependent cytotoxicity of Lipofectamine. This lipid, which forms DNA liposomes that fuse to cell membranes, offers several advantages such as high transfection efficiencies and targeting of the oligonucleotide to the nucleus (Bennett et al. 1992).

Unlike the nonspecific inhibitors, antisense oligonucleotides reduce the expression of a single protein product without disrupting other proteins responsible for normal cell function. Phosphorothioates are oligonucleotides with a sulfur substituted for one of the oxygens in the phosphodiester bonds between the nucleotides. This modification increases the stability of the oligonucleotides, especially in aqueous solutions, and prevents cellular nuclease enzymes from rapidly degrading them. Phosphorothioates can function in vivo by a number of mechanisms. At the RNA level they generate a substrate for RNase H activity by forming a thermally stable DNA-RNA duplex with a specific sequence and thus render the mRNA accessible for RNase-H cleavage (Stein 1998). In addition, antisense oligonucleotides can form triple helices that interfere directly with transcription and may also inhibit translation (Cazenave et al. 1989). In general, HPLC purified oligonucleotides ranging in size from 14 to 28 bases offer excellent specificity without being subject to self-annealing (Sczakiel1997).

Transient transfections

Although the insect steroid ponasterone A activates antisense expression, the decrease in binding may be attributed to either a low-level basal transcription of antisense DNA in the absence of ponasterone or, possibly, to Lipofectamine-mediated cell injury. It is notable that results using antisense transfections varied widely, possibly due to experimentally controlled differences in the number of cells that simultaneously internalize both plasmids and because of Lipofectamine’s transfection ratio relative to its toxicity. For instance, it was determined that a 4 h transfection maximized transfection efficiency while minimizing cytotoxicity. The plasmid’s effectiveness at reducing receptor expression in two of the three experiments may perhaps be due to the production of endogenously produced antisense mRNA, rather than a single transfection with exogenous oligonucleotides.

Unlike the use of exogenous antisense mRNA to downregulate GnRHR expression, transfection with a plasmid containing an insert encoding for a specific antisense sequence offers several advantages, including the ability to create a stable cell line containing the construct. This cell line would permit control over receptor-mediated processes by allowing suppression of receptor expression.

The ecdysone-inducible mammalian expression system utilizes a heterodimer of the ecdysone receptor (VgECR) and the retinoid X receptor (RXR) that binds a hybrid ecdysone response element (E/GRE) in the presence of the synthetic analogs of ecdysone, ponasterone A, or muristerone A. Ponasterone A is comparable to muristerone A in its affinity for the ecdysone receptor, induction of molting, and lack of endogenous effects in rodents, and is more readily available. The ecdysone receptor (VgEcR), modified to contain the VP16 transactivation domain, and the retinoid-X receptor (which is the mammalian homologue of USP, the natural partner to the ecdysone receptor) are both expressed from the pVgRXR vector. In the presence of ecdysone analogs, RXR and VgEcR associate together, bind to the hybrid response element, and activate transcription of the insert. This system allows for a tight regulation of expression in the absence of ponasterone A.

One reason why this initial study focused on GnRHR levels and not mRNA is the observation that GnRHR mRNA and protein levels are only sometimes, but not always, concordant (Sukhbaatar et al. 2015) (Sakurai et al. 1997). Thus, we chose, in this initial proof of concept study, to measure the final biological effect of our intervention and plan on conducting quantitative mRNA level measurements going forward. The authors are confident that the reduction in GnRHR is attributable to the antisense transcript produced by the inducible plasmid and not an artifact of transfection since the sense probe did not show an effect. The RXR plasmid is tightly regulated. Once a cell line was created, it was maintained in the OFF state and then split into two groups. The group receiving ponasterone A (a steroid derived from insects) showed a significant decrease in GnRHR expression when compared to cells untreated with this steroid and to controls. Cells from the OFF group were subsequently treated with ponasterone A and showed significant decrease in GnRHR (data not shown). As discussed above, cells transfected with the ‘sense’ insert, described previously, did not show any meaningful decrease in GnRHR expression even when incubated in the presence of ponasterone A (Figure 4). This allows us to conclude that the decrease in GnRHR expression is not due to an artifact of transfection but rather to the effect of the transcription of a specific antisense sequence.

Because experiments were conducted on different days, with different cell passage numbers and because the half-life of 125I-labeled GnRH agonist des-Gly10-[D-Ala6] GnRH N-ethylamide is 60 days, each experiment was run with its own control and data were normalized to controls.

Conclusion

GnRH-producing neurons originate from the olfactory placode, and after migration into the brain, are present in relatively low numbers in the hypothalamus. These cells function as the master regulators of the reproductive system and have the intrinsic ability to produce GnRH pulses. Clinically, a deeper understanding of GnRH pulse generation will help elucidate the molecular mechanisms behind the onset of puberty, better understand conditions such as idiopathic hypogonadotropic hypogonadism and certain forms of infertility. New medications capable of modulating GnRH production at a level upstream of the GnRH agonists and antagonists currently being prescribed can also be conceived. Thus, to study the mechanism underlying the pulsatile activity, we must first eliminate the autocrine effect of GnRH by downregulating the expression of the GnRHR.

GT1–7 GnRH-producing neurons are easily grown in vitro in the absence of external neural, endocrine, and autocrine inputs. This research has shown that it is possible to decrease receptor expression significantly using three modalities and that antisense techniques provide maximal decrease in GnRHR expression while minimizing cytotoxicity. It also suggests that the most effective method for downregulating the receptor would involve the generation of a stable cell line (instead of a transient transfection) or, possibly, a knockout construct. At this point, further work is required to test if the achieved reduction in GnRHR expression is sufficient to suppress the ultra-short feedback loop and alter GnRH production profiles.

Materials and methods

Cells

GT1–7 cells (generously provided by Dr. Richard Weiner, University of California, San Francisco) were grown in a 1:1 mixture of Dulbecco’s modified Eagle’s medium (DMEM) and F-12 medium (Gibco BRL, Grand Island, NY) with 2.5 g/L bicarbonate (Sigma Aldrich, St. Louis, MO) and 100 mg/L Gentamicin (Gibco) supplemented with 10% heat-inactivated fetal bovine serum (Biofluid, Port Richey, FL) (Mores et al. 1996). Cells were plated in 24-well culture plates (Costar, Corning, Corning, NY) at a density of 500,000 cells per well and incubated at 37°C under 5% CO2 in air.

Binding studies

Each well was washed three times with serum-free binding medium and cells were acid-washed prior to conducting binding studies. The binding medium consisted of DMEM (Sigma), 3.5 g/L Hepes (Sigma), 1 g/L BSA (Sigma), pH of 7.4. Acid washing of cells was performed using a solution containing 150 mM sodium chloride and 50 mM acetic acid to displace binding of endogenous GnRH.

Specific binding was determined by subtracting the nonspecific binding from the total binding. The latter was measured by adding 100,000 CPM per well of 125I-labeled GnRH agonist des-Gly10-[D-Ala6] GnRH N-ethylamide (Covance Laboratories, Madison, WI) corresponding to a concentration of 2 nM. Nonspecific binding was determined in the presence of 1 μM unlabeled GnRH analogue des-Gly10-[D-Ala6] GnRH N-ethylamide (Peninsula laboratories, San Carlos, CA), mixed with the radiolabeled agonist.

Cells were then incubated at room temperature for 45 min before removing all fluid from the wells and lysing the cells using 1 mL of 10 M NaOH and 10% sodium dodecyl sulfate. The lysate was transferred to a test tube and each well was subsequently rinsed with 1 mL of water prior to analysis in a gamma counter. Specific binding values were normalized to total protein, which was measured using the BCA Protein Assay Kit (Pierce Biotechnology, Rockford, IL).

Inhibition of RNA and protein synthesis

Cells were seeded in a 24-well plate (Corning) and incubated in culture medium for 24 h prior to a 5-day treatment with antineoplastic agents that suppress RNA or protein synthesis every 48 h. Antibiotics were added to regular culture medium to keep the following concentrations: 100 μg/mL puromycin; 10 μL/mL cycloheximide; 1 μg/mL actinomycin D and 50 μg/mL of α-amanitin, a specific inhibitor of RNA polymerase II (all from Sigma). Cell-surface GnRHR expression was assessed by binding studies and adjusted to total cellular protein content as described above, then normalized to 24 h controls.

Antisense oligonucleotides

Four HPLC-purified 21-mer antisense oligonucleotides (Gibco BRL) were obtained to selectively reduce expression of the GnRHR. The first of these phosphorothioates, AS-1 [5′-GTTACGCATATTTCTCCAAGG-3′], is complementary to a sequence spanning the start codon, while AS-2 [5′-AAGGAAGAAAGTCACGGTCAC-3′] targets a site located in the beginning of the GnRHR gene. A third sequence, ‘Sense’ [5′-AAGAATAATATCCCAAGAGCT-3′], which was picked at random, was used as a control to measure any nonspecific interaction between oligonucleotides and the transcription/translation of our gene of interest. AS-4 [5′-TCGCGTGAGAGCAAA-3′], located in the middle of the gene, was designed to optimize the Gibbs free energy (Branch 1998), while taking into account the secondary structure (Advanced Gene Computing Technologies, Irvine, CA).

GT1–7 cells (120,000) were plated in each well of a 24-well plate and incubated over a 48 h period in regular culture medium. Cells were transfected by replacing the medium with serum-free media containing 800 μL Opti-MEM I (Gibco) and 2.4 μL Lipofectamine reagent (Gibco), which resulted in a 400 nM oligonucleotide concentration. Every 48 h cells were transfected for a period of 4 h to reduce the effects of Lipofectamine toxicity and subsequently grown in regular medium to promote proliferation. After 5 days, the level of GnRHR expression was measured by binding studies. Data was adjusted to total protein content and normalized to 48 h controls.

Stable expression of antisense mRNA

To increase the intracellular concentration of antisense mRNA and further downregulate receptor expression, three different stable cell lines, each expressing a different inducible insert, were generated. Initially, a PCR reaction using the Elongase kit (Gibco) was used to amplify portions of our gene of interest from a plasmid containing the cloned full-length GnRHR gene. The first insert, S-1, which is close to the full length of the gene, was obtained using a forward primer (C) [5′-TGGAGAAATATGGCTAAC-3′] commencing about 10 bases prior to the start codon and an antisense primer (A) [5′-GTCTCCCAACTACAAAGA-3′] spanning the stop codon. The second insert, S-2, was constructed using the same forward primer and an antisense primer (T) [5′-CACCTTCATCCTTGAGA GCTT-3′] located about 243 bp downstream from the sense primer. The final insert, S-3 [5′-AAGGAA GAAAGTCACGGTCACATTGTTAGCCATATTTCTCCAAGGTAG-3′], was comprised of the fusion of the antisense oligonucleotides AS-1 and AS-2, which were described in the previous section, and included the addition of a stop codon. This latter sequence was chosen based on the potential success of the combination of oligonucleotides. Unlike the other two inserts, S-3 (as well as its complementary strand) were obtained using the Beckman Oligo 1000 DNA Synthesizer (Beckman Instruments, Fullerton, CA) and later annealed by incubation for 1 h at 65°C.

The products of the PCR reaction were run on 3% NuSieve agarose (FMC Bioproducts, Philadelphia, PA) and gel purified using the QIAquick Gel Extraction Protocol (Qiagen, Valencia, CA). A standard tailing reaction, using Taq DNA polymerase (promega, Madison, WI), was used to adenylate the blunt-end PCR fragments as well as the fusion oligonucleotide. The insert was cloned in one step into the pIND/V5-His-Topo vector (Invitrogen, San Diego, CA), a pre-linearized TA cloning plasmid containing the ampicillin and G418 resistance genes. Escherichia coli Top 10 Ultracomp cells (Invitrogen) were transformed by heat shock, following the One Shot protocol. After a 30 min incubation in SOC medium (Invitrogen), bacteria were plated onto 100 mm Petri dishes (BD Biosciences, Bedford, MA) containing Luria-Bertani Broth with 1.5% agar (KD Medical, Columbia, MD) supplemented with 50 μg/mL ampicillin.

Ampicillin-resistant colonies were transferred from the Petri dish into 50 mL Falcon Tubes (BD Biosciences, Bedford, MA) containing 5 mL Luria-Bertani Broth (Biofluids) supplemented with 50 μg/ mL ampicillin and incubated overnight in aerobic conditions in a shaker set at 37°C. Using the spin protocol for the Wizard Plus SV Miniprep kit (Promega), plasmid DNA was extracted for each bacterial colony. A PCR reaction, utilizing the Ecdysone forward primer [5′-CTCTGAATACTTTCAACAAGTTAC-3′] located 114 bases upstream of the multiple cloning site with the antisense primers A1 [5′-TGGAGAAATATGGCT AACAAT-3′] for the S2 insert or the A3 primer [5′-CTACCTTGGAGAAAT-3′] for the S3 fusion insert, was used to confirm the presence of an insert and to screen its orientation.

Colonies in which the insert was shown to be in the correct (antisense) orientation were inoculated into a flask containing 350 mL LB broth supplemented with ampicillin and grown overnight in a shaker set at 37°C. DNA was obtained using the Qiagen Maxi Protocol (Qiagen). The DNA was sequenced using the Thermo Sequenase (Amersham) dGTP protocol.

Prior to generating stable cell lines, transient cotransfections were performed using the RXR plasmid in concert with the insert containing the S-1 full-length antisense transcript of the GnRHR. Both plasmids, which were in the circularized DNA form, were transfected as described above for the phosphorothioate oligonucleotides. The insert was activated by adding ponasterone directly into the regular culture medium.

In order to generate stably transfected GT1–7 cells, the pIND plasmid DNA was linearized using the restriction enzyme PVU-1 (promega). The RXR plasmid, which was cotransfected into mammalian cells and permits tightly regulated expression of the insert, was linearized using the restriction enzyme MLU-1 (Promega). The DNA was subsequently washed using three volumes of ethanol and 30% ammonium acetate (both Sigma), rinsed with 70% ethanol, and redissolved in TE buffer.

GT1–7 cells were plated in a 24-well plate and allowed to grow to 80% confluency prior to transfection with 100 μL Opti-Mem I solution containing 1.5 μL Lipofectamine 2000 (Gibco) and 1 μg plasmid DNA. Forty-eight hours after co-transfection with RXR and the plasmid containing the insert, culture medium supplemented with 500 μg/mL Zeocin (Invitrogen) and 500 μg/mL Geneticin G418 (Gibco) was used to select double transfectants. Specific binding studies data was adjusted to total cellular protein and normalized to controls as previously described.

Statistical analysis

Statistical analysis was performed using SPSS (SPSS Inc. SPSS for Windows, Version 17.0. Chicago, IL). Data was analyzed by two-way analysis of the variance (ANOVA) in Figures 1 and 3 and by a oneway ANOVA in Figures 2 and 4, followed by Bonferroni post-hoc tests. p < 0.05 was considered significant.

Acknowledgments

This research was performed under a NIH-Intramural Research Training Award (IRTA). As a Medical Scholar in the Endocrinology and Reproductive Research Branch (NICHD-ERRB), I wish to thank my mentors Dr. Kevin J. Catt, Dr. Maria Dufau, and Dr. Lazar Krsmanovic. I am grateful for the excellent comments of an unknown reviewer. I also wish to thank the NIH-Women’s Reproductive Health Research Career Development Award (K-12) at Wayne State University for ongoing research support.

Abbreviations:

ANOVA

analysis of the variance

DMEM

Dulbecco’s modified Eagle’s medium

GnRH

gonadotropin-releasing hormone

RXR

retinoid X receptor

Footnotes

Disclosure statement

No potential conflict of interest was reported by the authors.

Institutional ethics approval

This project has no ethical impediments and consisted only of cell culture work using a commercially available cell line.

References

  1. Anderson SE, Dallal GE, Must A. 2003. Relative weight and race influence average age at menarche: results from two nationally representative surveys of US girls studied 25 years apart. Pediatrics. 111(4 Pt 1):844–850. [DOI] [PubMed] [Google Scholar]
  2. Austoker JL, Beebee TJ, Chesterton CJ, Butterworth PH. 1974. DNA-dependent RNA polymerase activity of Chinese hamster kidney cells sensitive to high concentrations of alpha-amanitin. Cell. 3(3):227–234. [DOI] [PubMed] [Google Scholar]
  3. Bedran de Castro JC, Khorram O, McCann SM. 1985. Possible negative ultra-short loop feedback of luteinizing hormone releasing hormone (LHRH) in the ovariectomized rat. Proc Soc Exp Biol Med. 179(1):132–135. [DOI] [PubMed] [Google Scholar]
  4. Bennett CF, Chiang MY, Chan H, Shoemaker JE, Mirabelli CK. 1992. Cationic lipids enhance cellular uptake and activity of phosphorothioate antisense oligonucleotides. Mol Pharmacol. 41(6):1023–1033. [PubMed] [Google Scholar]
  5. Bourguignon JP, Franchimont P. 1981. [Intermittent liberation of gonadoliberin (GnRH) by the rat hypothalamus in vitro]. C R Seances Soc Biol Fil. 175(3):389–396. [PubMed] [Google Scholar]
  6. Branch AD. 1998. A good antisense molecule is hard to find. Trends Biochem Sci. 23(2):45–50. [DOI] [PubMed] [Google Scholar]
  7. Cazenave C, Stein CA, Loreau N, Thuong NT, Neckers LM, Subasinghe C, Helene C, Cohen JS, Toulme JJ. 1989. Comparative inhibition of rabbit globin mRNA translation by modified antisense oligodeoxynucleotides. Nucleic Acids Res. 17(11):4255–4273. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Chason RJ, Kang JH, Gerkowicz SA, Dufau ML, Catt KJ, Segars JH. 2015. GnRH agonist reduces estrogen receptor dimerization in GT1–7 cells: evidence for cross-talk between membrane-initiated estrogen and GnRH signaling. Mol Cell Endocrinol. 404:67–74. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Chumlea WC, Schubert CM, Roche AF, Kulin HE, Lee PA, Himes JH, Sun SS. 2003. Age at menarche and racial comparisons in US girls. Pediatrics. 111(1):110–113. [DOI] [PubMed] [Google Scholar]
  10. Clarke H, Dhillo WS, Jayasena CN. 2015. Comprehensive review on Kisspeptin and its role in reproductive disorders. Endocrinol Metab (Seoul). 30(2):124–141. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Constantin S, Iremonger KJ, Herbison AE. 2013. In vivo recordings of GnRH neuron firing reveal heterogeneity and dependence upon GABAA receptor signaling. J Neurosci. 33(22):9394–9401. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Drouva SV, Laplante E, Kordon C. 1983. Effects of ovarian steroids on in vitro release of LHRH from mediobasal hypothalamus. Neuroendocrinology. 37(5):336–341. [DOI] [PubMed] [Google Scholar]
  13. Engmann L, DiLuigi A, Schmidt D, Nulsen J, Maier D, Benadiva C. 2008. The use of gonadotropiN-releasing hormone (GnRH) agonist to induce oocyte maturation after cotreatment with GnRH antagonist in high-risk patients undergoing in vitro fertilization prevents the risk of ovarian hyperstimulation syndrome: a prospective randomized controlled study. Fertil Steril. 89 (1):84–91. [DOI] [PubMed] [Google Scholar]
  14. Fritz MA, Speroff L. 2011. Clinical gynecologic endocrinology and infertility. 8th ed. Philadelphia, PA: Wolters Kluwer Health/Lippincott Williams & Wilkins. [Google Scholar]
  15. Herbison AE, Moenter SM. 2011. Depolarising and hyperpolarising actions of GABA(A) receptor activation on gonadotrophin-releasing hormone neurones: towards an emerging consensus. J Neuroendocrinol. 23(7):557–569. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. HermaN-Giddens ME, Slora EJ, Wasserman RC, Bourdony CJ, Bhapkar MV, Koch GG, Hasemeier CM. 1997. Secondary sexual characteristics and menses in young girls seen in office practice: a study from the pediatric research in office settings network. Pediatrics. 99(4):505–512. [DOI] [PubMed] [Google Scholar]
  17. Hu L, Gustofson RL, Feng H, Leung PK, Mores N, Krsmanovic LZ, Catt KJ. 2008. Converse regulatory functions of estrogen receptor-alpha and -beta subtypes expressed in hypothalamic gonadotropin-releasing hormone neurons. Mol Endocrinol. 22(10):2250–2259. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Humaidan P, Alsbjerg B. 2014. GnRHa trigger for final oocyte maturation: is HCG trigger history? Reprod Biomed Online. 29(3): 274–280. [DOI] [PubMed] [Google Scholar]
  19. Kanasaki H, Oride A, Mijiddorj T, Sukhbaatar U, Kyo S. 2017a. How is GnRH regulated in GnRH-producing neurons? Studies using GT1–7 cells as a GnRH-producing cell model. Gen Comp Endocrinol. 247:138–142. [DOI] [PubMed] [Google Scholar]
  20. Kanasaki H, Tselmeg M, Oride A, Sukhbaatar U, Hara T, Kyo S. 2017b. Pulsatile kisspeptin effectively stimulates gonadotropin-releasing hormone (GnRH)-producing neurons. Gynecol Endocrinol. 33(9):721–727. [DOI] [PubMed] [Google Scholar]
  21. Kedinger C, Gniazdowski M, Mandel JL Jr., Gissinger F, Chambon P. 1970. Alpha-amanitin: a specific inhibitor of one of two DNA-pendent RNA polymerase activities from calf thymus. Biochem Biophys Res Commun. 38(1):165–171. [DOI] [PubMed] [Google Scholar]
  22. Knobil E. 1981. Patterns of hypophysiotropic signals and gonadotropin secretion in the rhesus monkey. BiolReprod. 24(1):44–49. [DOI] [PubMed] [Google Scholar]
  23. Knobil E. 1990. The GnRH pulse generator. Am J Obstet Gynecol. 163(5 Pt 2):1721–1727. [DOI] [PubMed] [Google Scholar]
  24. Krsmanovic LZ, Martinez-Fuentes AJ, Arora KK, Mores N, Navarro CE, Chen HC, Stojilkovic SS, Catt KJ. 1999. Autocrine regulation of gonadotropiN-releasing hormone secretion in cultured hypothalamic neurons. Endocrinology. 140(3):1423–1431. [DOI] [PubMed] [Google Scholar]
  25. Krsmanovic LZ, Stojilkovic SS, Balla T, al-Damluji S, Weiner RI, Catt KJ. 1991. Receptors and neurosecretory actions of endothelin in hypothalamic neurons. Proc Natl Acad Sci U S A. 88(24):11124–11128. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Krsmanovic LZ, Stojilkovic SS, Merelli F, Dufour SM, Virmani MA, Catt KJ. 1992. Calcium signaling and episodic secretion of gonadotropin-releasing hormone in hypothalamic neurons. Proc Natl Acad Sci U S A. 89 (18):8462–8466. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Larco DO, Williams M, Schmidt L, Sabel N, Lange J, Woller MJ, Wu TJ. 2015. Autoshortloop feedback regulation of pulsatile gonadotropin-releasing hormone (GnRH) secretion by its metabolite, GnRH-(1–5). Endocrine. 49(2):470–478. [DOI] [PubMed] [Google Scholar]
  28. Levy MN, Koeppen BM, Stanton BA. 2005. Principles of physiology. New York: Mosby. [Google Scholar]
  29. Liposits Z, Merchenthaler I, Wetsel WC, Reid JJ, Mellon PL, Weiner RI, Negro-Vilar A. 1991. Morphological characterization of immortalized hypothalamic neurons synthesizing luteinizing hormone-releasing hormone. Endocrinology. 129(3):1575–1583. [DOI] [PubMed] [Google Scholar]
  30. Maggi R, Cariboni AM, Marelli MM, Moretti RM, Andre V, Marzagalli M, Limonta P. 2016. GnRH and GnRH receptors in the pathophysiology of the human female reproductive system. Hum Reprod Update. 22:3. [DOI] [PubMed] [Google Scholar]
  31. Mellon PL, Windle JJ, Goldsmith PC, Padula CA, Roberts JL, Weiner RI. 1990. Immortalization of hypothalamic GnRH neurons by genetically targeted tumorigenesis. Neuron. 5 (1):1–10. [DOI] [PubMed] [Google Scholar]
  32. Moenter SM, Caraty A, Lehman MN, Karsch FJ. 1993. Characterization and regulation of pre-ovulatory secretion of gonadotrophin-releasing hormone. Hum Reprod. 8 (Suppl 2):51–56. [DOI] [PubMed] [Google Scholar]
  33. Moghissi KS. 2000. A clinician’s guide to the use of gonadotropin-releasing hormone analogues in women. Medscape Womens Health. 5(1):5. [PubMed] [Google Scholar]
  34. Mores N, Krsmanovic LZ, Catt KJ. 1996. Activation of LH receptors expressed in GnRH neurons stimulates cyclic AMP production and inhibits pulsatile neuropeptide release. Endocrinology. 137(12):5731–5734. [DOI] [PubMed] [Google Scholar]
  35. Nunez L, Faught WJ, Frawley LS. 1998. Episodic gonadotropiN-releasing hormone gene expression revealed by dynamic monitoring of luciferase reporter activity in single, living neurons. Proc Natl Acad Sci U S A. 95 (16):9648–9653. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Rogers MC, Silverman AJ, Gibson MJ. 1998. Preoptic area grafts implanted in mammillary bodies of hypogonadal mice: patterns of GnRH neuronal projections. Exp Neurol. 151(2):265–272. [DOI] [PubMed] [Google Scholar]
  37. Roth C, Schricker M, Lakomek M, Strege A, Heiden I, Luft H, Munzel U, Wuttke W, Jarry H. 2001. Autoregulation of the gonadotropin-releasing hormone (GnRH) system during puberty: effects of antagonistic versus agonistic GnRH analogs in a female rat model. J Endocrinol. 169(2):361–371. [DOI] [PubMed] [Google Scholar]
  38. Sakurai H, Adams BM, Adams TE. 1997. Concentration of GnRH receptor and GnRH receptor mRNA in pituitary tissue of orchidectomized sheep: effect of oestradiol, progesterone, and progesterone withdrawal. J Endocrinol. 152(1):91–98. [DOI] [PubMed] [Google Scholar]
  39. Sczakiel G. 1997. The design of antisense RNA. Antisense Nucleic Acid Drug Dev. 7(4):439–444. [DOI] [PubMed] [Google Scholar]
  40. Silverman A. 1994. The gonadotropin releasing hormone (GnRH), neuronal systems: immunocyto-chemistry and in situ hybridization. New York: The Physiology of Reproduction Raven Press Ltd; p. 1683–1709. [Google Scholar]
  41. Stein CA. 1998. How to design an antisense oligodeoxynucleotide experiment: a consensus approach. Antisense Nucleic Acid Drug Dev. 8(2):129–132. [DOI] [PubMed] [Google Scholar]
  42. Stojilkovic SS, Krsmanovic LZ, Spergel DJ, Catt KJ. 1994. Gonadotropin-releasing hormone neurons: intrinsic pulsatility and receptor-mediated regulation. Trends Endocrinol Metab. 5(5):201–209. [DOI] [PubMed] [Google Scholar]
  43. Sukhbaatar U, Kanasaki H, Mijiddorj T, Oride A, Miyazaki K. 2013. Kisspeptin induces expression of gonadotropin-releasing hormone receptor in GnRH-producing GT1–7 cells overexpressing G proteiN-coupled receptor 54. Gen Comp Endocrinol. 194:94–101. [DOI] [PubMed] [Google Scholar]
  44. Sukhbaatar U, Mijiddorj T, Oride A, Kanasaki H. 2015. Stimulation of delta subunit-containing GABAA receptor by DS1 increases GnRH receptor expression but reduces GnRH mRNA expression in GnRH-producing GT1–7 cells. Endocrine. 49(1):222–230. [DOI] [PubMed] [Google Scholar]
  45. Van Goor F, Krsmanovic LZ, Catt KJ, Stojilkovic SS. 2000. Autocrine regulation of calcium influx and gonadotropin-releasing hormone secretion in hypothalamic neurons. Biochem Cell Biol. 78(3):359–370. [PubMed] [Google Scholar]
  46. Weiner RI W, Goldsmith P, Martinez de la Escalera G, Windle J, Padula C, Choi A, Negro-Vilar A, Mellon P. 1992. Gonadotropin-releasing hormone neuronal cell lines. Front Neuroendocrinol. 13(2):95–119. [PubMed] [Google Scholar]
  47. Wetsel WC. 1995. Immortalized hypothalamic luteinizing hormone-releasing hormone (LHRH) neurons: a new tool for dissecting the molecular and cellular basis of LHRH physiology. Cell Mol Neurobiol. 15(1):43–78. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Wetsel WC, Valenca MM, Merchenthaler I, Liposits Z, Lopez FJ, Weiner RI, Mellon PL, Negro-Vilar A. 1992. Intrinsic pulsatile secretory activity of immortalized luteinizing hormone-releasing hormone-secreting neurons. Proc Natl Acad Sci U S A. 89(9):4149–4153. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Woller M, Tessmer S, Neff D, Nguema AA, Roo BV, Waechter-Brulla D. 2001. Leptin stimulates gonadotropin releasing hormone release from cultured intact hemihypothalami and enzymatically dispersed neurons. Exp Biol Med (Maywood). 226(6):591–596. [DOI] [PubMed] [Google Scholar]
  50. Woller MJ, Meyer S, Ada-Nguema A, Waechter-Brulla D. 2004. Dissecting autocrine effects on pulsatile release of gonadotropin-releasing hormone in cultured rat hypothalamic tissue. Exp Biol Med (Maywood). 229(1):56–64. [DOI] [PubMed] [Google Scholar]
  51. Wray S. 2010. From nose to brain: development of gonadotrophin-releasing hormone-1 neurones. J Neuroendocrinol. 22(7):743–753. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Wray S, Hoffman G. 1986. A developmental study of the quantitative distribution of LHRH neurons within the central nervous system of postnatal male and female rats. J Comp Neurol. 252(4):522–531. [DOI] [PubMed] [Google Scholar]
  53. Wu TJ, Mani SK, Glucksman MJ, Roberts JL. 2005. Stimulation of luteinizing hormone-releasing hormone (LHRH) gene expression in GT1–7 cells by its metabolite, LHRH-(1–5). Endocrinology. 146(1):280–286. [DOI] [PubMed] [Google Scholar]

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