Cloning and functional characterization of the ovine Hormone Sensitive Lipase (HSL) full-length cDNAs: An integrated approach

Gene 416 (2008) 30–43

Contents lists available at ScienceDirect

Gene j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / g e n e

Cloning and functional characterization of the ovine Hormone Sensitive Lipase (HSL) full-length cDNAs: An integrated approach Antonis D. Lampidonis a, Alexandros Argyrokastritis a,1, Dimitrios J. Stravopodis b, Gerassimos E. Voutsinas c, Triantafyllia G. Ntouroupi d, Lukas H. Margaritis b, Iosif Bizelis a,⁎, Emmanuel Rogdakis a a

Department of Animal Science, Laboratory of Animal Breeding and Husbandry, Agricultural University of Athens, Iera Odos 75, 118 55 Athens, Greece Department of Cell Biology and Biophysics, Faculty of Biology, University of Athens, Panepistimiopolis, Zografou 157 84, Athens, Greece Institute of Biology, National Center for Scientific Research (NCSR) “Demokritos”, Agia Paraskevi 153 10, Athens, Greece d Weatherall Institute of Molecular Medicine, University of Oxford, John Radcliffe Hospital, Headington, Oxford, OX3 9DS, UK b c

A R T I C L E

I N F O

Article history: Received 18 September 2007 Received in revised form 20 February 2008 Accepted 21 February 2008 Available online 12 March 2008 Received by M. D'Urso Keywords: Adipose tissue Cloning Hormone Sensitive Lipase (HSL) Ovis aries PCR

A B S T R A C T Hormone Sensitive Lipase (HSL) is a highly regulated enzyme that mediates lipolysis in adipocytes. HSL enzymatic activity is increased by adrenergic agonists, such as catecholamines and glucagons, which induce cyclic AMP (cAMP) intracellular production, subsequently followed by the activation of Protein Kinase A (PKA) and its downstream signalling cascade reactions. Since HSL constitutes the key enzyme in the regulation of lipid stores and the only enzyme being subjected to hormonal regulation [in terms of the recently identified Adipose Triglyceride Lipase (ATGL)], the ovine Hormone Sensitive Lipase (ovHSL) fulllength cDNA clones were isolated, using a Polymerase Chain Reaction-based (PCR) strategy. The two isolated isoforms ovHSL-A and ovHSL-B contain two highly homologous Open Reading Frame (ORF) regions of 2.089 Kb and 2.086 Kb, respectively, the latter having been missed the 688th triplet coding for glutamine (ΔQ688). The putative 695 and 694 amino acid respective sequences bear strong homologies with other HSL protein family members. Southern blotting analysis revealed that HSL is represented as a single copy gene in the ovine genome, while Reverse Transcription-PCR (RT-PCR) approaches unambiguously dictated its variable transcriptional expression profile in the different tissues examined. Interestingly, as undoubtedly corroborated by both RT-PCR and Western blotting analysis, ovHSL gene expression is notably enhanced in the adipose tissue during the fasting period, when lipolysis is highly increased in ruminant species. Based on the crystal structure of an Archaeoglobus fulgidus enzyme, a three-dimensional (3D) molecular model of the ovHSL putative catalytic domain was constructed, thus providing an inchoative insight into understanding the enzymatic activity and functional regulation mechanisms of the ruminant HSL gene product(s). © 2008 Elsevier B.V. All rights reserved.

1. Introduction Every organism, including ruminants, continuously disburses energy not only for movement or to perform labor, but also for maintenance, regulation and almost every other biological process. Orga-

⁎ Corresponding author. E-mail address: [email protected] (I. Bizelis). Abbreviations: ATGL, Adipose Triglyceride Lipase; BFAE, Brefeldin A Esterase; BSA, Bovine Serum Albumin; cAMP, Cyclic AMP; CHO, Chinese Hamster Ovaries; DTT, Dithiothreitol; FA, Fatty Acid(s); GAPDH, Glyceraldehyde-3-Phosphate Dehydrogenase; HSL, Hormone Sensitive Lipase; ORF, Open Reading Frame; ovHSL, Ovine HSL; PCR, Polymerase Chain Reaction; PDB, Protein Data Bank; PKA, Protein Kinase A; PMSF, Phenyl-Methyl-Sulfonyl-Fluoride; PNPB, p-Nitrophenyl Butyrate; RACE, Rapid Amplification of cDNA End(s); RT-PCR, Reverse Transcription-PCR; STH, Strand-Turn-Helix; TAG(s), Triacylglycerol(s); UTR(s), Untranslated Region(s) (5′- / 3′-); WAT, White Adipose Tissue; 3D, Three-Dimensional. 1 The present work is dedicated to the memory of Dr. Alexandros Argyrokastritis, Lecturer, who, so suddenly, passed away by heart attack, on the 22nd of August 2005.

0378-1119/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.gene.2008.02.026

nisms derive their energy from the breakdown of nutrients they take up. The type of nutrients can roughly be divided in three groups: proteins, carbohydrates and lipids. During breakdown of these nutrients, energy is produced in the form of ATP, which is the universal energy carrier in all organisms. Most nutrients, however, are not directly utilized for generation of energy, but are rather stored in tissues in an appropriate form for later occasions. Lipids are mainly stored as triacylglycerol (TAG) in the adipose tissues and comprise essential components of ruminant diets for three reasons, given as follows: (a) The high caloric content of lipids can be a valuable factor for overcoming limitations in energy supplies in high yielding ruminants; (b) Lipids can be used to modulate the digestion and absorption profile of different nutrients. For example, fats can constrain ruminant acidosis and depressed milk fat content, resulting from high carbohydrate and low fiber diets (Palmquist, 1984). Moreover, lipid intake can alter the proportion of particular fatty acids (FA) in meat or milk fat, to be more desirable for the food industry or for consumption by humans (Gnunmer, 1991); (c) Some vegetable or

A.D. Lampidonis et al. / Gene 416 (2008) 30–43

animal lipids constitute low priced feedstuffs that may be of particular interest in the formulation process of ruminant diets. During periods of nutrient deprivation and/or stress, a complex metabolic process, called lipolysis, is carried out by adipocytes, in which non-esterified fatty acids and glycerol are liberated from the triacylglycerol storage droplets and released from the cell. Being an important process for the regulation of energy homeostasis, lipolysis is implemented by at least three adipocyte-specific enzymes. A monoglyceride lipase catalyzes the breakdown of monoglycerides to glycerol and fatty acids, whereas a Hormone Sensitive Lipase, so-called HSL, mediates for the hydrolysis of triglycerides to diglycerides and diglycerides to monoglycerides (Holm, 2003). Besides, a novel lipase termed Adipose Triglyceride Lipase (ATGL), alternatively denoted as either desnutrin or iPLA2 has been recently identified (Jenkins et al., 2004; Villena et al., 2004; Zimmermann et al., 2004). Using antibodies directed against ATGL lipase, Zimmerman et al. (2004) proposed that ATGL is responsible for the 75% of total cytosolic acylhyrolase activity in white adipose tissue (WAT) of HSL deficient mice. ATGL could, therefore, participate together with HSL, in a likely cooperative fashion, for the efficient implementation of the WAT lipolysis process. Using a highly specific HSL inhibitor, the relative contribution of HSL and ATGL respective lipolytic activities has been recently re-evaluated (Langin et al., 2005). By subsuming all the available observations, a putative biochemical partitioning of the cellular lipase activity could be successfully adopted for human fat cells, as following: Even though both ATGL and HSL enzymes possess strong capacities to efficiently hydrolyze triglycerides moieties in vitro (Fredrikson et al., 1981; Zimmermann et al., 2004), only HSL bears a significant diglyceride lipase activity (Haemmerle et al., 2002), also being the only enzyme tightly subjected to hormonal regulation (Holm, 2003). HSL [gene symbol LIPE, EC 3.1.1.3 (Kraemer and Shen, 2002)] constitutes a highly regulated enzyme that mediates lipolysis in adipocytes. One of the most important inducers of lipolysis is the catecholamine family members, which can significantly increase cyclic AMP (cAMP) intracellular levels, subsequently followed by the activation of Protein Kinase A (PKA) and its signalling cascade network (Holm, 2003). HSL lipolytic activity is substantially augmented by β-adrenergic agonists and glucagon, whereas insulin evinces an opposite effect, causing a significant reduction in HSL enzymatic potency (Holm et al., 2000). After β-adrenergic stimulation, HSL is phosphorylated at several serine residues by cAMP-dependent PKA (Holm et al., 2000) and an extracellular signal-regulated kinase (Greenberg et al., 2001). The phosphorylated HSL is transported from the cytoplasm to the surface of a lipid droplet (Egan et al., 1992). Conversely, a lipid droplet surface protein also phosphorylated by PKA, so-called perilipin A, shifts to the cytoplasm in response to lipolytic stimulation (Brasaemle et al., 2000). Moreover, it has been previously reported that HSL also binds to the docking protein lipotransin (Syu and Saltiel, 1999) and to a fatty acid-binding protein (Shen et al., 1999). HSL is expressed at low levels in a variety of different cell lines and tissues (Holm et al., 1987; Holm et al., 1988), including heart (Small et al., 1989), skeletal muscle (Langfort et al., 1998), macrophages (Jepson and Yeaman, 1993; Contreras et al., 1994), pancreatic β-cells (Klannemark et al., 1998) and cultured Chinese Hamster Ovaries (CHO) cells (Osuga et al., 1997). The enzyme aggregates at high concentration, requires the presence of detergents for purification and binds to phospholipid vesicles, properties suggestive of a hydrophobic protein (Holm et al., 1990). In the present study, we describe the isolation and functional characterization of the full-length cDNA clones, encoding the two ovHSL protein isoforms A and B. The cloning strategy was based on an RT-PCR approach, combined by RACE-PCR reactions applied on both 5′- and 3′-ends. As it is unambiguously revealed by Southern blotting analysis, the ovHSL-A and ovHSL-B proteins are produced by a single copy gene and likely represent two allelic or polymorphic derivatives

31

[without excluding alternative mechanisms (i.e. differential splicing, or editing)], whereat ovHSL-B is distinguished from ovHSL-A by the absence of only one amino acid residue (ΔQ688). Howbeit its variable transcriptional activation profile, ovine HSL gene expression is notably induced in adipose cells during fasting conditions, a valuable observation that is clearly corroborated by both RT-PCR and Western blotting approaches. Multiple sequence alignments, escorted by molecular modeling design, were able to dictate the presence of evolutionary conserved structural elements, which could likely play essential roles in the catalytic activity and hormonal regulation courses of the ruminant HSL proteins. 2. Materials and methods 2.1. Total RNA purification from ovine adipose tissue and Reverse Transcription (RT) reactions Total RNA was isolated from ovine adipose tissue according to the manufacturer's protocol of “RNeasy Lipid Tissue Kit” (Qiagen, USA). First-strand cDNA synthesis was carried out with Invitrogen “Thermoscript RT-PCR System”, using 1 μg of purified total RNA. Briefly, total RNA preparations together with 50 μM of oligo-[dT]20, 10 mM of dNTP mix and DEPC-treated water were denatured at 65 °C for 5 min. After adding cDNA synthesis buffer, 100 mM DTT, 40 U RNaseOUT and 15 U of reverse transcriptase up to 20 μl of total reaction volume, an annealing and polymerization step at 60 °C for 1 h was carried out. The reaction was terminated by incubating at 85 °C for 5 min. 2.2. Isolation of cDNA clones encoding ovine homologues of HSL using Polymerase Chain Reaction (PCR) PCR amplification reactions were carried out using deoxynucleotides, buffers and enzyme (Taq polymerase) concentrations as exactly recommended by the enzyme manufacturer (New England Biolabs, USA). Reactions were performed on a GeneAmp PCR System 2400 thermocycler with an initial denaturation step at 94 °C for 3 min, followed by 40 cycles at 94 °C for 30 s, 60 °C for 30 s and 72 °C for 2.5 min. A polymerization step at 72 °C for 10 min was added after the completion of 40 cycles. The amplification reactions were carried out using the pre-synthesized ovine cDNA as a template (under standard PCR conditions) and the appropriate set of primers were designed from highly conserved regions of sequence identity among the published human, mouse, rat, pig and bovine sequences [i.e. Bt5_for / HSL2_rev set of primers were used in a typical PCR reaction] (Table 1; Fig. 1). 2.3. cDNA cloning of the 5′- and 3′-untranslated regions through RACE Rapid Amplification of cDNA end(s) (RACE) is a PCR-based technique, which facilitates the cloning of full-length cDNA sequences when only a partial cDNA fragment is available. According to this Table 1 Nomenclature, nucleotide sequences and corresponding length of all the primers utilized for the molecular cloning and isolation of the ovHSL cDNAs Name

Oligonucleotide sequence (5′ → 3′)

Bt5_for HSL2_rev Bt5_rev Bt5N_rev 5′-RACE inner 3′-RACE inner HSL3_for HSL4_for GAPDH_for GAPDH_rev HSL1_for HSL1_rev

CTACCTGGCCGCCCTCACC GAAGGAGTTGAGCCACGAGG GTCTCGTTGCGTTTGTAGTGC GCAGCGGCCGTAGAAGCAG CGCGGATCCGAACACTGCGTTTGCTGGCTTTGATG CGCGGATCCGAATTAATACGACTCACTATAGG GTGCTTCTACGCCTACTGCTG TCCGACTCAGACCAGAAGGC TGGTATCGTGGAAGGACTCATGAC ATGCCAGTGAGCTTCCCGTTCAGC CCTACCTCAAGAACTGGGCC CTGGGTAGGCTGCCATGATG

Length 19 20 21 19 35 32 21 20 24 24 20 20

nt nt nt nt nt nt nt nt nt nt nt nt

32

A.D. Lampidonis et al. / Gene 416 (2008) 30–43

Fig. 1. Graphical presentation of the cloning strategy designed for the ovHSL cDNAs isolation. Bt5_for and HSL2_rev primers, synthesized according to a phylogenetically conserved region originated from the bovine HSL cDNA sequence, were utilized in RT-PCR reactions, resulting in the production of a 1.25 Kb ovine cDNA homologous family member. Through 5′-RACE-PCR and 3′-RACE-PCR approaches, three additional cDNA fragments were cloned, with notated length 0.615 Kb (5′-RACE inner / Bt5N_rev set of primers), 0.955 Kb (HSL3_for / 3′-RACE inner set of primers) and 0.71 Kb (HSL4_for / 3′-RACE inner set of primers). Based on all possible partial overlaps among the four clones (0.615 Kb, 1.25 Kb, 0.955 Kb and 0.71 Kb), a linearly contiguous assembly of the obtained ovHSL cDNAs was generated and two respective full-length genetic forms (ovHSL-A and ovHSL-B) were recognized (see Fig. 3). The HSL1_for / HSL1_rev set of primers (0.27 Kb) was used for the RT-PCR mediated assessment of ovHSL gene expression profile (see Fig. 6A), while the 0.11 Kb cDNA fragment produced by the HSL4_for / HSL2_rev set of primers was utilized in a Southern blotting-mediated examination of ovHSL gene copy number in the ovine genome (see Fig. 5).

principle and using the FirstChoice RLM-RACE Kit (Ambion, USA), the missing sequences of the ovHSL cDNA 5′- and 3′-untranslated regions [UTR(s)] were targeted for cloning. Reverse Transcription procedure, 5′-RACE-PCR outer and 5′-RACEPCR inner reactions were carried out using 1 μg of total purified RNA and Taq Polymerase (New England Biolabs, USA), according to the manufacture's protocol (Ambion, USA). The amplification reactions were performed using as gene-specific primers the Bt5_rev (outer) and Bt5N_rev (inner) ones, having, alongside, a distance of 0.1 Kb (Table 1; Fig. 1). Reactions were carried out on a GeneAmp PCR System 2400 thermocycler, with an initial denaturation step at 94 °C for 3 min, followed by 35 cycles at 94 °C for 30 s, 60 °C for 30 s and 72 °C for 3 min. A polymerization step at 72 °C for 7 min was added after the completion of 35 cycles. The molecular cloning of the ovHSL cDNA 3′-UTR was attained according to the Invitrogen (USA) “Thermoscript RT-PCR System”. First-strand cDNA synthesis was carried out using 1 μg of the 3′-RACE adaptor (FirstChoice RLM-RACE Kit-Ambion, USA). The amplification reactions were performed using as gene-specific primers the HSL3_for (outer) and HSL4_for (inner) ones (Table 1; Fig. 1), according to the FirstChoice RLM-RACE Kit-Ambion (USA). Reactions were carried out on a GeneAmp PCR System 2400 thermocycler, following the same conditions described above. 2.4. Molecular cloning and DNA sequencing analysis of the ovHSL PCR fragments The obtained PCR fragments were gel purified using the Qiaquick Gel Extraction Kit (Qiagen, USA), ligated to vector pGEM-T-Easy (Promega, USA), transformed into Escherichia coli JM109 competent cells and plated on appropriate indicator LB-dishes containing ampicillin as a selection reagent. Plasmids carrying the various ovHSL cDNA PCR fragments were isolated using the Qiaprep Miniprep Kit (Qiagen, USA) and three independent clones of each fragment were sequenced three times in both directions (Institute of Molecular Biology and Biotechnology, Foundation of Research and Technology, Heraklion, Crete, Greece).

enzymes AvaII, EcoRI, NcoI and PstI. After separation of the obtained genomic fragments through electrophoresis, their DNA denaturation and subsequent transfer from the agarose gel to a nitrocellulose membrane, a Southern hybridization process was carried out to determine the informative genomic restriction fragments length, using as a probe a [32P]-labeled 0.11 Kb RT-PCR fragment of the ovHSLA cDNA, derived from the HSL4_for / HSL2_rev set of primers (Table 1; Fig. 1), near the 3′-coding end. Hybridization reactions were performed overnight at 65 °C under high stringency conditions, as described by Sambrook et al. (1989). 2.6. Semi-quantitative Reverse Transcription (RT)-PCR approach for the assessment of ovHSL gene expression profile First-strand cDNA synthesis was carried out with the Qiagen “Omniscript Reverse Transcription” kit (Qiagen, USA), using mRNA preparations isolated from ovine adipose tissue during (a) the refeeding period of the sheep, which lasted two weeks and (b) the fasting period with also two weeks durance, the latter having provided 25% of the required energy, as well as from muscle, adrenal gland and testis tissue under normal diet conditions. The RT-PCR reactions were quantitatively analyzed using as a comparative control product a part of the coding region of the ovine GAPDH (Glyceraldehyde-3-Phosphate Dehydrogenase) gene, amplified with the specific forward primer GAPDH_for and the reverse GAPDH_rev one (Table 1), whereas the condition (refeeding or fasting)-dependent and tissuespecific ovHSL gene expression profile was examined through the utilization of the specific set of internal primers HSL1_for and HSL1_rev (Table 1; Fig. 1). All PCR reactions were performed using equal corresponding amounts of templates and primers in each one of them, while 30 cycles for either ovHSL or GAPDH amplified fragments were run in order to ensure the long-linear range of the reactions. The PCR scheduled conditions for product amplifications are given as following: 94 °C for 3 min, ensued by 30 cycles of 94 °C for 30 s, 60 °C for 30 s and 72 °C for 1 min, with a final extension step at 72 °C for 7 min (Taq polymerase, New England Biolabs, USA). 2.7. Western blotting analysis and ovHSL protein immunodetection

2.5. Southern blotting analysis examining the ovHSL gene copy number Genomic DNA from ovine blood was isolated using the GFX Genomic Blood DNA Purification Kit (Amersham Biosciences, USA) and subsequently digested (15 μg / reaction) with the restriction

Frozen adipose tissue samples (dissected during refeeding or fasting periods) with an approximate mass of 100 mg each were thoroughly squashed in a liquid nitrogen containing pot and subsequently homogenized in a solubilisation lysis buffer (pH 7.6, at 4 °C),

A.D. Lampidonis et al. / Gene 416 (2008) 30–43

containing 1% Triton X-100, 50 mM NaCl, 30 mM Na4P2O7 (× 10H2O), 50 mM NaF, 0.1 mM Na3VO4 (× H2O), 5 mM Na2EDTA (× 2H2O), 0.1% BSA (Bovine Serum Albumin), 50 mM Tris-base and 2 mM PMSF (Phenyl-Methyl-Sulfonyl-Fluoride), the latter reagent being utilized as a strong and irreversible protease inhibitor (added fresh each time). The cellular homogenates were centrifuged at 12,500 rpm for 30 min in 4 °C. The obtained supernatant was re-diluted in an 1:1 molecular ratio with 2× Laemmli buffer [100 mM Tris-HCl pH 6.8, 200 mM DTT (Dithiothreitol), 4% SDS, 0.2% Bromophenol Blue and 20% Glycerol] and finally centrifuged at 12,500 rpm for 15 min in 4 °C, targeting to the sequestration of the insoluble material to the pellet and the acquisition of all the protein components in the aqueous supernatant phase. A 10% SDS-PAGE denaturing gel electrophoresis was performed, as described by Laemmli (1970), on a BIORAD (USA) miniprotean apparatus. The separated proteins were electro-transferred onto a nitrocellulose membrane (Amersham-Pharmacia, USA), using as transfer-buffer an aqueous solution containing 25 mM Tris-base, 200 mM glycine and 20% methanol. The ovHSL protein was immunodetected using a rabbit polyclonal antibody (Cat. No. 10006371, Cayman, USA), at a final working dilution of 1:400 in blocking buffer (1 × TBS-T and 5% non-fat milk), before a secondary antibody comprised by an anti-rabbit IgG conjugate (Cat. No. A4914, SigmaAldrich, USA) was utilized in a dilution of 1:2000 under the same incubation conditions (1 × TBS-T and 5% non-fat milk). Finally, the ovHSL immobilized protein on the nitrocellulose membrane was visualized using the ECL system, according to manufacturer's instructions (Amersham-Pharmacia, USA). In order to be able to quantitatively validate the obtained ovHSL protein-band densities, with regard the total protein amount of our cellular extracts, a standard anti-α-tubulin monoclonal antibody (Cat. No. 2125 S, Cell Signalling, USA) was utilized in a final dilution of 1:3000 in blocking buffer (1× TBS-T and 5% BSA), a process being directly followed by the incubation with an anti-mouse IgG conjugate (Cat. No. A6782, Sigma-Aldrich, USA) in an 1:2000 working dilution (1× TBS-T and 5% non-fat milk) and finally completed by an ECL-mediated protein-band visualization.

33

Residues 265 to 687 of the predicted ovHSL polypeptide sequence were appropriately modeled using as a suitable template the 1jji_A available structure of the, most highly homologous, hyper-thermophilic carboxylesterase from the archaeon Archaeoglobus fulgidus (De Simone et al., 2001) and a three-dimensional (3D) molecular model of the putative ovHSL protein was accordingly begot. 3. Results 3.1. Isolation and structural characterization of the ovHSL full-length cDNA clones RT-PCR reactions were performed on mRNA purified from ovine adipose tissue. A set of primers for PCR reactions was designed and synthesized according to the consensus regions derived through the complete nucleotide alignment of known bovine, human, mouse, rat and pig HSL cDNA sequences. Via the utilization of suitable software programs for primer design, two novel cDNA primers were generated, now called Bt5_for and HSL2_rev (Table 1; Fig. 1). The produced PCR fragment of the ~ 1.25 Kb expected size was initially resolved in a 1.5% agarose gel electrophoresis (Fig. 2A). The purified ~1.25 Kb cDNA moiety was inserted into the pGEM-T-Easy vector and three independent clones were completely sequenced three separate times from both strands, in order to ensure the reliability and fidelity of the obtained sequence information. Subsequent BLAST analysis of the ovine originating cDNA nucleotide sequence revealed an approximate 85% grade of identity with human, mouse, rat and pig respective cDNA regions, whereas bovine and ovine seems to share a 95% of HSL cDNA sequence identity.

2.8. In silico analysis and molecular modeling of ovHSL protein The primers design was carried out using the OligoAnalyser v.1.2 and OligoExplorer v.1.2 software system (http://molbiol-tools. ca/molecular_biology_freeware.htm). The functional analysis of the sequenced clones was performed using a conventional BLAST search procedure (http://www.ncbi.nlm.nih.gov/BLAST/), with the support of online supplementary programs (http://biology.semo.edu/cgi-bin/ dnatools.pl). Sequence alignments for the detection of distinct structural domains and functional motifs throughout ovHSL putative protein were implemented via the utilization of the multiple alignment software package Multalin (http://prodes.toulouse.inra.fr/multalin/ multalin.html). The generation of a reliable phylogenetic tree of the HSL family members and their molecular evolution course was attained through the comparison of the putative ovHSL protein(s) primary structure(s) (present study) with homologous amino acid sequences from other model organisms, such as Bos indicus (bovine), Homo sapiens (human), Mus musculus (mouse), Rattus norvegicus (rat), Sus scrofa (pig) and Ciona intestinalis (Ciona-sea squirt). Protein domains were identified in the putative ovHSL amino acid sequence through its comparison to related sequences in the Pfam database (http://www.sanger.ac.uk/Software/Pfam/). Secondary structure predictions of the ovHSL putative protein were performed by the PSIPRED Protein Structure Prediction Server [http://bioinf.cs.ucl.ac.uk/ psipred/], exactly as previously described (Jones, 1999). The ovHSL protein was modeled by means of the Swiss-Model and SwissPdbViewer molecular graphics modeling packages (http://swissmodel. expasy.org/), according to the similarities of the modeled sequence to known structures, publicly available in the Protein Data Bank (PDB).

Fig. 2. RT-PCR-mediated cloning and purification of the ovHSL cDNA fragments, whose contiguous assembly results in the generation of full-length clones. (A) Amplification of a cDNA fragment of 1.25 Kb (lane F), using the flanking primers Bt5_for / HSL2_rev. (B) Amplification of a cDNA fragment of 0.615 Kb (lane Q), using the flanking primers Bt5N_rev / 5′-RACE inner, containing the 5′-UTR, through a 5′-RACE-PCR approach. (C) Amplification of a cDNA fragment of 0.955 Kb (lane S) and one of 0.710 Kb (lane R), using the flanking primers HSL3_for / 3′-RACE inner and HSL4_for / 3′-RACE inner, respectively, containing the 3′-UTR, through a 3′-RACE-PCR approach. Numbers on the left indicate the molecular sizes of the marker bands lane (MW).

34

A.D. Lampidonis et al. / Gene 416 (2008) 30–43

The characterization of the ovHSL cDNA missing segments, including the 5′- and 3′-untranslated regions (UTRs) was ensued by a RACEPCR approach, using as a primary template total RNA purified from ovine adipose tissue. A 5′-RACE-PCR reaction on the already isolated cDNA clone of ~ 1.25 Kb, via the utilization of the Bt5N_rev and 5′RACE inner primers (Table 1; Fig. 1), gave rise to the amplification of a 5′-allocated 0.615 Kb fragment (Fig. 2B). Similarly, in the context of a 3′-RACE-PCR molecular strategy and through the utilization of two set of primers HSL3_for / 3′-RACE inner and HSL4_for / 3′-RACE inner (Table 1; Fig. 1), two 3′-allocated 0.955 Kb and 0.71 Kb cDNA fragments were respectively produced (Fig. 2C). After the insertion of the purified 5′- and 3′-cDNA moieties into the pGEM-T-Easy host vector, three independent clones were completely sequenced three separate times each in both strand-orientations. As expected, all three (0.615, 0.955 and 0.71 Kb) RACE-PCR products, after the appropriate sequence alignments, bore 100% nucleotide identity with the respective overlapping regions of the ~1.25 Kb fragment. Interestingly, in contrast to 0.71 Kb, the 0.955 Kb 3′-RACE-PCR product revealed to contain an additional triplet coding for glutamine (Q) and an extra segment of 24 bp at the proximal non-coding 3′-UTR. Thus, the two distinct cDNAs of 2.248 Kb (ovHSL-A) and 2.221 Kb (ovHSL-B) seems to retain two almost identical ORFs of 2.089 Kb and 2.086 Kb respectively, which consequently represent the two ovHSL putative isoforms. The structural characteristics that can distinguish the ovHSL-A cDNA from the ovHSL-B one, are pin-pointed in the insertion of an extra glutamine-coding triplet at the corresponding protein position 688 (Q688) and the incorporation of a 24 bp nucleotide portion at the proximal 3′-UTR (Fig. 3). To avoid any technical artifact presumably

derived from the 3′-RACE-PCR approach, three independent recombinant clones were separately sequenced three times from both strands, thus ensuring DNA sequencing validity and accuracy. 3.2. Molecular dissection of ovHSL cDNAs and proteomic annotation of their putative isoforms The re-constructed full-length nucleotide sequences, containing the corresponding ORFs, as well as their flanking 3′- and 5′-UTRs, of both ovHSL-A and ovHSL-B cDNA clones were appropriately deposited in the GenBank database obtaining the accession numbers DQ647325 and DQ647326, respectively. The ovHSL-A full-length cDNA clone is characterized by a coding region of 2.089 Κb, a 5′- and 3′-UTR comprised by 72 bp and 100 bp, respectively, the latter being followed by a poly-[A] tail region (Fig. 3A). On the contrary, the ovHSL-B fulllength cDNA clone contains a coding region of 2.086 Κb, flanked by a 5′-UTR of 72 bp and a 3′-UTR of 76 bp (instead of 100 bp, as occurs for ovHSL-A) adjoining with a poly-[A] tail area (Fig. 3B). The nucleotide sequence surrounding the translation initiation codon AUG corresponding position, on both ovHSL-A and ovHSL-B cDNAs, 5′-AGGatgG-3′ stands in absolute conformity with the Kozak vertebrate consensus sequence [A/G]XXAUGG (Kozak, 1981). Both cDNA clones seem to contain an identical 5′-UTR sequence that is mainly characterized by the presence of an out of frame stop codon (UGA), thus rendering the 5′-border of the translation starting site an abstruse and inaccurate trait. A crucial structural feature unambiguously distinguishing the ovHSL-A from its counterpart allelic isoform ovHSL-B is pin-pointed in a 3′-UTR located

Fig. 3. Graphical presentation and sequence decipherment of the two ovHSL full-length cDNA clones and their putative proteins. (A) The ovHSL-A cDNA, comprised of 2.248 Kb, contains the whole coding area, along with the flanking 5′- and 3′-untranslated regions (UTRs). Note the translation initiation codon ATG and the stop codon TAA. The 5′-UTR and 3′UTR are denoted by dash lines, while the asterisks represent the two canonical poly-adenylation signals located 5′-upstream (47 bp and 35 bp, respectively) of the poly-[A] tail. The full-length ovHSL-A cDNA includes both the CAG triplet coding for the Q688 residue and the 24 bp 3′-UTR sequence “TGGGCTTGACTCATCCTGAAATCG”. (B) The ovHSL-B cDNA, comprised of 2.221 Kb, contains the whole coding area, along with the flanking 5′-UTR and 3′-UTR. Note the translation initiation codon ATG and the stop codon TAA. The 5′-UTR and 3′-UTR are denoted by dash lines, while the asterisks represent the two canonical poly-adenylation signals located 5′-upstream (23 bp and 11 bp, respectively) of the poly-[A] tail. The ovHSL-B cDNA is characterized by the absence of (a) the CAG triplet coding for the glutamine residue at position 688 (ΔQ688) and (b) the 24 bp 3′-UTR sequence denoted as Δ24. (C) Nucleotide sequence of the ovHSL-A full-length cDNA clone and its cognate putative protein of 695 amino acid residues. Note the Kozak sequence (indicated by a box) surrounding the ATG (bold fonts) translation initiation codon and the stop codon TAA, along with a second in frame stop codon TGA (to ensure the termination of translation), both denoted by bold fonts. The two canonical poly-adenylation signals AATAAA are underlined, whereas the poly-[A] tail is shown by italic fonts. The, likely, allelic or polymorphic differences of ΔQ688 and Δ24 (Β) in between the ovHSL-A and ovHSL-B cDNAs are both indicated by grey coloring.

A.D. Lampidonis et al. / Gene 416 (2008) 30–43

35

Fig. 3 (continued ).

region of 24 bp that has been missing in the case of ovHSL-B cDNA clone. Moreover, the 3′-UTR of both cDNA clones seems to contain two canonical poly-adenylation signals (AAUAAA) each, which are

located 47 bp and 35 bp 5′-upstream of the ovHSL-A, as well as 23 bp and 11 bp 5′-upstream of the ovHSL-B poly-[A] tail areas, respectively (Fig. 3C).

36

A.D. Lampidonis et al. / Gene 416 (2008) 30–43

Regarding the molecular anatomy of the putative ovHSL protein isoforms, the ovHSL-A ORF is comprised by a 695 amino acid sequence, resulting in a predicted MW of 76.5 kDa and a theoretical pI of 6.27, whereas the ovHSL-B ORF proclaims for the occurrence of an almost identical to the ovHSL-A protein isoform of 694 amino acid residues, predicating for a MW of 76.4 kDa and a pI of 6.27. Interestingly, as it is clearly demonstrated in Fig. 4, the putative 695 (ovHSL-A) and 694 (ovHSL-B) amino acid respective sequences

bear strong homologies with other HSL protein family members, an observation that strongly vindicates both their evolutionarily conserved and physiologically essential characters in a variety of different species (Fig. 9). More specifically, an elaborated BLAST analysis and a multiple alignment approach revealed that both ovHSL-A and ovHSLB protein isoforms manifest a significantly high degree of amino acid sequence identity with: (a) B. indicus (bovine) protein (88%), (b) H. sapiens (human) protein (76%), (c) M. musculus (mouse) protein (74%),

Fig. 4. Multiple amino acid sequence alignment of ovHSL-A and ovHSL-B proteins together with representative HSL protein family members. Ovine (Ovis A and Ovis B) (Ovis aries: accession number of ovHSL-A {Ovis A}: DQ647325; accession number of ovHSL-B {Ovis B}: DQ647326), bovine (Bos indicus: NM_001080220), human (Homo sapiens: NM_005357), mouse (Mus musculus: NM_001039507), rat (Rattus norvegicus: NM_012859) and pig (Sus scrofa: NM_214315). Notice (a) the short COOH-terminal hepta-peptide tail “GSPTSGS” that is characterized by a S / T rich motif and (b) the critical Q (glutamine residue) at position 688, clearly distinguishing the ovHSL-A (695 amino acid residues) protein isoform from its ovHSL-B (694 amino acid residues) highly homologous counterpart.

A.D. Lampidonis et al. / Gene 416 (2008) 30–43

37

(d) R. norvegicus (rat) protein (76%) and (e) S. scrofa (pig) protein (79%) (Fig. 4). Interestingly, the comparative multi-protein identity pattern seems to include the putative starting methionine, thus unambiguously corroborating the functional role of its corresponding AUG as the veritable initiation codon of the translation process (Fig. 3). Examination of the structural composition of both ovHSL-A and ovHSL-B amino acid sequences, targeting for the recognition of functionally important domains, revealed that a highly homologous, among all family members, NH2-terminal domain is observed between positions 1–315. A distinct and catalytically fundamental region could be designated between the amino acid residues 333 and 499 of the ovHSL isoforms, mainly being characterized by the significantly high degree of amino acid sequence identity (92%) with the rat HSL homologous protein. Interestingly, this middle region appears to contain the active site serine residue (S424), thus likely bearing an enzymatic activity against the water soluble p-Nitrophenyl Butyrate (PNPB) substrate (Tsujita et al., 1989). Moreover, it is mainly characterized by the presence of a highly conserved, among almost all lipases, GXSXG motif that is located at the positions 422–426 of both ovHSL isoforms. Smith et al. (1996), as well as Holm et al. (1994), have previously suggested that this is the bona fide catalytic domain of HSL, as they have demonstrated that substitution of serine 424 (S424) directly results to the complete abolition of esterase and lipase activity. The COOH-terminal region of a representative HSL protein family member contains two serine phosphorylation sites (residues 563 and 565 of rat HSL) that seem to be conserved and can be undoubtedly recognized in the ovHSL isoforms A and B at positions 552 (S552) and 554 (S554). Shen et al. (1998) have previously demonstrated that S552 is phosphorylated by cyclic AMP-dependent protein kinase (PKA), whereas S554 is phosphorylated by Ca2+/calmodulin-dependent protein kinase II. The phosphorylation and, moreover, the subsequent activation of ovHSL proteins are likely implemented inside the COOHterminal regulatory region that is located between the amino acid residues 499–647 of both ovHSL isoforms (Smith et al., 1996). Finally, a distinct structural domain so-called α/β hydrolase fold, which represents a centrally arranged β sheet surrounded by a variable number of α helices, is located between 346 and 550 (V346–R550) amino acid residues. 3.3. Assessment of ovHSL gene copy number and its rough structural organization, as evinced by a Southern blotting approach Total genomic DNA, purified from ovine blood was digested with different restriction endonucleases (AvaII, PstI, NcoI and EcoRI) and subsequently treated, following the Southern blotting protocol of Sambrook et al. (1989). After genomic band separation in a 1% agarose gel, DNA blotting and high stringency hybridization reaction, through the utilization of a 0.11 Kb RT-PCR cDNA fragment, located close to the ovHSL-A 3′-coding end, single bands corresponding to the AvaII and PstI restriction fragments and two major hybridization zones reflecting the NcoI and EcoRI generated genomic moieties were easily observed even in the short term exposures of the related autoradiographs (Fig. 5). The obtained pattern of ovHSL genomic structure and the previous knowledge that the homologous gene family members seem to contain a large number of various sizes introns (Li et al., 1994; Langin et al., 1993; Harbitz et al., 1999) constitute major arguments allowing us to strongly suggest that HSL is likely represented as a single copy gene locus in the ovine genome. 3.4. Tissue-specific expression profile of the ovHSL gene: a fasting condition induced response A tissue-specific pattern of the ovHSL transcriptional activity was obtained after total RNA isolation from ovine adipose tissue during refeeding and fasting periods, as well as from muscle, adrenal gland and testis tissue, subsequently followed by an RT-PCR approach. The

Fig. 5. Detection of the HSL gene in the ovine genome, as evinced by a Southern blotting approach. (A) Ovine genomic DNA, after its purification, was digested with the restriction endonucleases NcoI, AvaII, EcoRI and PstI (lanes) and subsequently separated in 1% agarose gel electrophoresis. The obtained pattern was visualized through Ethidium-Bromide staining under a UV-lamp (312 nm). (B) Autoradiograph of the agarose gel described in (A), after its treatment through a Southern blotting procedure and final hybridization under high stringency conditions, using as a probe a [32P]labelled RT-PCR fragment of 0.11 Kb, generated close to the 3′-coding end of the ovHSLA cDNA clone, using the HSL4_for / HSL2_rev set of primers.

0.27 Kb (Fig. 6A) generated product likely reflects the ovHSL gene transcriptional activity, without excluding alternative regulatory mechanisms, such as an mRNA stabilization process. As it is clearly illustrated in Fig. 6A and B, the ovHSL gene is variably expressed among the examined ovine tissues, but most interestingly is notably induced (~ 2×) in adipose tissue during the fasting period of the sheep (Fig. 6B). Due to the variable levels of GAPDH gene activity among the examined tissues, as clearly evinced through the production of a 0.189 Kb RT-PCR fragment (Fig. 6A) and in order to more accurately estimate the ovHSL tissue-dependent transcript accumulation, the obtained band intensities for each ovHSL and GAPDH fragment were converted to a number, through utilization of the Scion-Image software tool (Scion Corporation, USA). Subsequently, a numeral ratio of ovHSL / GAPDH was formulated, thus allowing a more reliable assessment of the ovHSL transcriptional expression profile in each of the examined tissues (Fig. 6B). Interestingly, in all the RT-PCR set of performed reactions (n = 5), the relative transcript abundance of ovHSL (i.e. ovHSL / GAPDH) was noticeably enriched in adipose tissue and, moreover, repeatedly induced of approximately 2× under fasting diet

38

A.D. Lampidonis et al. / Gene 416 (2008) 30–43

conditions (Fig. 6B). Statistical differences between the respective obtained values were analyzed using Student's t-test and the ones with P b 0.01 (Testis versus Adrenal gland; Muscle versus Adrenal gland) and P b 0.001 [Adipose tissue R (Refeeding) versus Adipose tissue F (Fasting); Adipose tissue R (Refeeding) versus Muscle; Adipose tissue R (Refeeding) versus Adrenal gland; Adipose tissue R (Refeeding) versus Testis; Testis versus Muscle] were considered statistically significant. In an effort to distinguish the potential differential expression of ovHSL-A and ovHSL-B isoforms during fasting conditions (Fig. 6), a transcript specific reverse primer was designed on the Δ24 3′-UTR region bearing the nucleotide sequence “5′-CGATTTCAGGATGAGTCAAGCC-3′”. Despite our strenuous efforts to specifically determine the ovHSL-A in vivo expression profile, through an RT-PCR approach, no cDNA fragment, using a variety of forward primers, could ever be detectably produced under all the examined conditions. This could be likely attributed to the intrinsic inability of the particular primer to efficiently anneal to its complementary sequence, albeit a prevailing representation of the second isoform ovHSL-B could not be excluded.

3.5. Fasting-dependent ovHSL protein accumulation in ovine adipose tissue Western blotting analysis using a rabbit anti-HSL polyclonal antibody (raised against rat HSL) resulted in the identification of a 76 kDa protein band (Fig. 7A) that was notably induced (~ 2×) in the adipose tissue during the fasting period of the sheep (Fig. 7B), thus corroborating the transcription expression profile of the ovHSL gene obtained under similar diet conditions (Fig. 6). Moreover, the p76ovHSL protein seems to be in terms of antigenicity similar to the rat HSL enzyme, as this has been rationally foreseen by the strong amino acid sequence homology of representative family members denoted in Fig. 4. Interestingly, the predicted molecular weight of the putative ovHSL proteins, as resulted from their cognate cDNA “coding calculations” (ovHSL-A: 75,6 and ovHSL-B: 75,4), strongly agrees with the in vivo estimated molecular weight of 76 kDa, thus strongly indicating that both ovHSL cDNAs constitute full-length clones containing the whole and complete Open Reading Frames (ORFs), respectively.

Fig. 6. Examination of the ovHSL gene expression profile through a semi-quantitative RT-PCR approach. (A) 250 ng of total RNA from each ovine tissue (lanes) were subjected to multiplex RT-PCR amplification reactions (30 cycles) and 25 μl of the RT-PCR product for each sample were separated in 2% agarose gel electrophoresis, whereat the obtained bands were visualized through Ethidium-Bromide staining under a UV-lamp (312 nm). To ensure the equal quantitation of the utilized templates, GAPDH was used as a reference gene. The production of the 0.27 Kb (primers: HSL1_for / HSL1_rev) and 0.189 Kb (primers: GAPDH_for / GAPDH_rev) RT-PCR fragments likely reflects the transcriptional activity of their cognate genes ovHSL and GAPDH, respectively. Notice the tissue-specific variable amplification pattern of the 0.27 Kb generated band and the latter's fasting-dependent induction in adipose tissue. (B) ovHSL relative transcript abundance, given as a numeral ratio of ovHSL / GAPDH band optical density, in each tissue examined in Fig. 6A. The normalized data are presented as mean values ± SEM (respective bars) of five separate experiments. ⁎⁎⁎: Statistically significant differences of relative transcript abundance (P b 0.001) between the respective pairs described in text. +++: Statistically significant differences of relative transcript abundance (P b 0.01) between Testis and Adrenal gland, as well as Muscle and Adrenal gland.

A.D. Lampidonis et al. / Gene 416 (2008) 30–43

Finally, the visualization of a 55 kDa protein zone, corresponding to the endogenous α-tubulin in both refeeding and fasting conditions (Fig. 7A), unambiguously demonstrates the structural integrity of our adipose tissue extracts, albeit a degree of quantitative fluctuation is observed. Therefore, the optical density of each 76 and 55 kDa band was accordingly measured (Scion-Image software tool, Scion Corporation, USA) and a numeral ratio of ovHSL / α-tubulin was produced, thus resulting to a more reliable assessment of the ovHSL protein representation in the examined adipose tissue extracts. As it is clearly illustrated in Fig. 7B, the relative ovHSL protein abundance (i.e. ovHSL / α-tubulin) in adipose tissue is undoubtedly increased in an approximate level of 2 × during the fasting period of the sheep, in all the performed experimental trials (n = 5), strongly corroborating the respective obtained results of the RT-PCR approach (Fig. 6). Statistical differences between the obtained data were analyzed according to Student's t-test and values P b 0.001 [Adipose tissue R (Refeeding) versus Adipose tissue F (Fasting)] were considered statistically significant.

39

3.6. Protein molecular modeling of the ovHSL catalytic domain A structural prediction of the ovHSL alpha / beta hydrolase fold, essentially constituting the catalytic domain, was constructed from the primary structure of the putative polypeptide containing the amino acid residues 265–687, through the utilization of the automated protein modeling server Swiss-model (Guex and Peitsch, 1997). Fig. 8 [(A)–(C)] clearly illustrates the six different views of the threedimensional (3D) structural model of the ovHSL enzyme, showing eight beta sheets connected by alpha helices, which literally represent the core structural elements of the alpha / beta hydrolase fold catalytic domain (Ollis et al., 1992). This structural unit, usually bearing diverse functional features and phylogenetic origins, is mainly comprised by a catalytic triad, whose elements are born on loops. As evinced by an in silico analysis (Fig. 4) and a molecular modeling approach, the serine residue at position 424 (S424) is predicated to reside into the core active site (Fig. 8D), likely modulating the alpha / beta hydrolase fold molecular conformation.

Fig. 7. Western blotting-mediated immunodetection of a fasting-induced accumulation of ovHSL endogenous protein in adipose tissue. (A) Whole protein cellular extracts were subjected to a 10% SDS-PAGE denaturing gel electrophoresis and subsequently transferred onto a nitrocellulose membrane. Incubation of the membrane with a rabbit anti-HSL polyclonal antibody resulted in the generation of a 76 kDa zone directly reflecting the endogenous levels of ovHSL protein(s) (upper panel). Notice the markedly induced p76ovHSL protein accumulation under fasting conditions in the adipose tissue. The structural integrity of protein extract components and their comparative quantitation, through preparation and loading procedures, were examined via the utilization of an anti-α-tubulin monoclonal antibody. The 55 kDa obtained zone represents the endogenous levels of α-tubulin that is expressed and accumulated in the ovine adipose tissue under refeeding and fasting conditions (bottom panel). (B) ovHSL relative protein abundance, given as a numeral ratio of ovHSL / α-tubulin band density, in the two tissues examined in Fig. 7A. The normalized data are presented as mean values ± SEM (respective bars) of five separate experiments. ⁎⁎⁎: Statistically significant differences of relative protein abundance (P b 0.001) between Adipose tissue R (Refeeding) and Adipose tissue F (Fasting).

40

A.D. Lampidonis et al. / Gene 416 (2008) 30–43

Due to severe limitations of protein structures availability deposited in Public Data Banks, the 3D molecular configuration of the ovHSL catalytic domain was constrainedly predicated through the utilization of the, most highly homologous [33% identity, against the other significant hit (31%) of Bacillus subtilis Brefeldin A Esterase (BFAE), whose crystal structure has been also resolved and deposited in PDB with a reference entry 1jkm (Wei et al., 1999)], hyperthermophilic carboxylesterase originated from A. fulgidus (De Simone et al., 2001). 4. Discussion Though ATGL catalyzes the rate-limiting step, Hormone Sensitive Lipase (HSL) plays an important role in the hydrolysis of stored triacylglycerols (TAGs) and is, therefore, a key enzyme in lipid metabolism and overall energy homeostasis (Fredrikson et al., 1981; Stralfors et al., 1987). As HSL has been extensively studied in human and other mammalian species, complete and annotated gene sequences have been already deciphered for human (Langin et al., 1993; Holst et al., 1996), mouse (Li et al., 1994; Sztrolovics et al., 1997), rat (Fredrikson et al., 1981; Østerlund et al., 1996), pig (Harbitz et al., 1999) and bovine (Cordle et al., 1986; Garton et al., 1988) species. Levitt et al. (1995) have mapped the HSL gene onto chromosomes 19p13.1–q13.2 regarding human, whereas, as it has been previously demonstrated by Warden et al. (1993), the mouse homologue is localized onto chromosome 7. According to the literature, no information concerning the ovHSL chromosomal topology has been reported so far. The molecular cloning and isolation of the ovine HSL (ovHSL) cDNAs were performed using an RT-PCR-based approach. The generation of linearly assembled cDNA fragments resulted in two almost identical ovHSL full-length clones, the ovHSL-A of 2.248 Kb and the ovHSL-B of 2.221 Kb. Multiple nucleotide sequence alignments with full-length cDNAs originated from different mammalian family members revealed strong identities within previously characterized areas. In regard with the cDNA untranslated regions, the ovine 5′-UTR and 3′-UTR seem to be quite shorter in comparison to the ones of other mammalian species. Therefore, we focused our efforts to clone and isolate longer transcripts using a variety of different set of primers. Even though we examined several experimental conditions, no alternative transcript could be ever isolated, thus strongly suggesting that the full-length ovHSL cDNAs truly contain surprisingly short 5′-UTR and 3′-UTR, in contrast to what has been observed in other mammalian species. Amino acid sequence alignment among the ovHSL putative protein and the mature family members of bovine, human, mouse, rat, pig and Ciona species was utilized as a major bio-informatic tool for generating a phylogenetic relationships tree, clearly illustrated in Fig. 9. ovHSL protein bears strong similarities with its bovine counterpart (Fig. 4), thus constituting the ruminant subgroup within the mammalian group of HSL proteins. The highly significant degree of primary structure conservation among representative vertebrate and invertebrate HSL protein family members presumably reflects the functional constrains during evolution, as they have been specifically adapted to fulfill each species individualized requirements. The occurrence of the ovine HSL homologous gene was unambiguously documented through a Southern blotting approach, clearly illustrated in Fig. 5. The obtained hybridization pattern of the ovHSL genomic locus and the large number of variably sized introns characterizing other HSL gene family members strongly indicate that, most probably, the HSL is represented as a single copy gene in the haploid version of sheep genome. Since the ovHSL-A and ovHSL-B full-length cDNA clones seem to encode two almost identical putative proteins, we further attempted to investigate the expression profile of the HSL gene in various ovine tissues, including the adipose one. Due to the inconclusive results obtained after the application of a Northern blotting approach, RT-PCR reactions were performed on total

Fig. 8. Molecular modeling of the ovHSL putative catalytic domain. (A)–(C) Six different views of the three-dimensional (3D) theoretical model of the ovHSL putative catalytic domain that contains the 265–687 amino acid residues, showing eight beta sheets connected by alpha helices, which literally represent the core structural elements of the alpha / beta hydrolase fold catalytic domain. (D) The ovHSL S424 (Ser424, depicted in yellow), along with the catalytic triad residues of the Archaeoglobus fulgidus hyperthermophilic carboxylesterase [S160 (Ser160), D255 (Asp255) and H285 (His285), depicted in red] and of the Brefeldin A Esterase (BFAE) [S202 (Ser202), D308 (Asp308) and H338 (His338), depicted in white], are illustrated in higher proportion. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

RNA preparations, revealing that ovHSL gene was variably expressed in the ovine tissues examined (Fig. 6A and B). Interestingly, one of the novel findings of the present study constitutes the fasting period-dependent induction of the ovHSL gene expression in the adipose tissue (Fig. 6B). During the fasting period of the sheep, lipolysis, characterized by an increased rate, leads to net loss of triglycerides from fat cells, thus generating the required energy for the metabolic demands of the organism. Similar results have been previously reported for human, rat and bovine HSL homologous gene family members (Holm et al., 1988; Small et al., 1989; Langfort et al., 2003; Martín-Hidalgo et al., 2005).

A.D. Lampidonis et al. / Gene 416 (2008) 30–43

In order to examine an implicit fasting condition-mediated ovHSL protein response, a Western blotting approach was applied and a 76 kDa protein was detected. The observed fasting period-dependent accumulation of the ovHSL protein in adipose tissue (Fig. 7A and B), strongly corroborating the RT-PCR obtained data (Fig. 6A and B), likely reveals the biological significance of the ovHSL enzymatic activity under “stress-genic” (fasting) conditions. Even though the fasting response is characterized by a notable ovHSL protein accumulation in adipose tissue, a process likely regulated by a transcriptional activation mechanism (Fig. 6), without excluding a protein stabilization scenario, no conclusive information can be obtained regarding the ovHSL phosphorylation status under different diet conditions. In order to reliably dissect the molecular anatomy of the ovHSL putative protein, multiple amino acid sequence alignments, containing the bovine, human, mouse, rat, pig and Ciona homologous family members, were performed. Bio-informatic analysis led to the identification of two distinct domains: a Hormone Sensitive Lipase NH2terminal (HSL_N) domain residing from 1-315 amino acid residues and an alpha / beta hydrolase fold domain (Abhydrolase_3) located between 346 and 550 residues. Moreover, another domain that could likely bear an enzymatic activity against the water soluble p-Nitro phenyl Butyrate (PNPB) synthetic substrate is recognized between 333 and 499 amino acid residues, thus strongly suggesting that it contains the active site serine residue (S424) and must be considered as the catalytic domain of ovHSL protein. Limited tryptic digestion of native HSL was previously demonstrated to dramatically reduce the activity of the enzyme against trioleoylglycerol substrate, whilst the activity against PNPB was relatively trypsin-insensitive (Smith et al., 1996). A distinct essential region, likely functioning as a regulatory domain, seems to reside between 499 and 647 ovHSL amino acid residues, also containing two putative phosphorylation sites pin-pointed at positions S552 and S554 (Shen et al., 1998). Interestingly, the functional effects of these critical serine residues could be quite different in that phosphorylation of the first (regulatory) site could activate the enzyme, whilst phosphorylation at the second (basal) site might not be able to induce any significant effect on activity and, moreover, could likely inhibit the subsequent phosphorylation at the regulatory site (Garton et al., 1989). As it was previously demonstrated by Anthonsen et al. (1998), the rat HSL protein contains two additional phosphorylation sites that are critically involved in the regulation of lipase activity. It has been undoubtedly shown that HSL protein is phosphorylated at serine residues 659 and 660, which are absolutely required for the phosphorylation-

41

induced increase in hydrolytic activity towards a triacylglycerol substrate, as well as for the translocation of HSL to the lipid droplet (Anthonsen et al., 1998; Su et al., 2003), upon over-expression of mutated at serine residues 563 and 565 HSL protein, along with an activated protein kinase A (PKA) isoform in cultured adipocytes. Interestingly, based on the multiple alignment of amino acid sequences illustrated in Fig. 4, only the second phosphorylation site of serine 649 can be unambiguously recognized in the ovHSL putative protein sequence, thus allowing us to reasonably speculate that S649 critically regulates ovHSL cellular activities. A unique anatomical feature of both ovHSL-A and ovHSL-B protein isoforms, clearly distinguishing them from the rest of the family members denoted in Fig. 4, represents the short COOH-terminal tail of “GSPTSGS” residues located at the corresponding positions 689–695. This hepta-peptide motif could be presumably phosphorylated on certain serine (S) and / or threonine (T) residues, so as to appropriately regulate lipase activity. Another novel and interesting finding of the present study is pinpointed in the, likely, allelic or polymorphic difference of ΔQ688 occurring in between the ovHSL-A and ovHSL-B enzyme isoforms. Even though alternative scenarios (i.e. alternative splicing or RNA editing) could not be excluded and since the Southern blotting analysis demonstrated the presence of a single copy gene, the absence of a glutamine (Q) residue at position 688 in the ovHSL-B putative protein is likely attributed to an allelic difference or polymorphic variation flowing through the ovine population examined in the present study. The missing part of 24 bp (“TGGGCTTGACTCATCCTGAAATCG”) (3′-UTR of ovHSL-B cDNA), located in between the second poly-adenylation signal and the poly-[A] tail, likely represents the second major allelic or polymorphic difference of the ovHSL genetic locus (the scenario of an alternative splicing process is still effective), whose functional importance remains to be determined. Focusing to the structural features of the ovHSL putative proteins, we generated in silico a three-dimensional (3D) molecular model of the 265–687 ovHSL-A / -B common protein segment. Since there does not seem to exist any resolved eukaryotic protein structure available in PDB, for applying it as a structural template platform, a theoretical model of the ovHSL alpha / beta hydrolase fold catalytic domain was constructed, through the utilization as a suitable scaffold of the crystal structure of the, most highly homologous, hyper-thermophilic carboxylesterase originated from the archaeon A. fulgidus (Fig. 8). Based on the multiple amino acid sequence alignment (Fig. 4), combined with a molecular modeling approach (Fig. 8), we strongly suggest that serine 424 (S424) occupies a central position in the active

Fig. 9. Molecular evolution of the ovHSL enzyme isoforms from a common ancestor. An evolutionarily distance-based and neighbour-joining phylogenetic tree, relating the ovHSL putative protein sequences to their family homologous member ones originated from bovine, pig, mouse, rat, human and Ciona species, was constructed through the utilization of PHYLIP program. A bootstrap analysis (151, 231 and 250 successful replicates) was performed using SEQBOOT. Distances were calculated via PROTDIST; the phylogenetic tree was subsequently generated by using NEIGHBOR and CONSENSE and drawn through the utilization of TREEVIEW. The programs SEQBOOT, PROTDIST, NEIGHBOR and CONSENSE are parts of PHYLIP program. Notice the generation of a ruminant subgroup, containing the Ovine and Bovine HSL protein family members.

42

A.D. Lampidonis et al. / Gene 416 (2008) 30–43

site and constitutes a critical residue of the catalytic triad. S424 lies in the middle of the conserved sequence GXSXG (aGDSAGg in the HSL protein family members, including ovHSL), which has been previously observed in a variety of different enzymes harboring the catalytic triad (Brenner, 1988). For reasons of molecular geometry and stereochemical constraints, the super-secondary structure Strand-Turn-Helix (STH) requires the occurrence of glycine residues (G) at positions +2 and −2 with respect to the nucleophile S424. Moreover, to appropriately shape the overall structure, a residue with a short lateral chain, alanine 425 (A425) in the case of HSL family members (Fig. 4), is placed after the critical serine. Fig. 8D illustrates the three-dimensional (3D) position of S424 (Ser424, depicted in yellow) in ovHSL protein, along with the respective catalytic triad residues of the A. fulgidus hyper-thermophilic carboxylesterase (depicted in red) and of the B. subtilis Brefeldin A Esterase (BFAE) (depicted in white). Intriguingly, the other two structural elements of the catalytic triad in human HSL, D693 (Asp693) and H723 (His723) (Contreras et al., 1996; Wei et al., 1999), are not respectively represented in the 265–687 ovHSL protein segment and, moreover, are located outside of the common homologous region among the examined family members (Fig. 4). Since the D693 residue in human HSL protein is replaced by a threonine [T692(A) / 691(B)] in sheep and the respective H723 residue seems to be missing from the ovHSL proteins, due to their shorter COOH-terminal tails, it is reasonable to speculate that the catalytic triad in the ovine member is conformed by S424 along with two additional amino acid residues that still remain elusive and need to be further explored. Finally, the molecular cloning and functional characterization of the ovHSL cDNA family members reveal novel and valuable primary information, thus allowing us to extrapolate reliable conclusions regarding their tissue-specific expression and diet condition-dependent regulation in ruminants. Acknowledgement The present work was kindly and generously supported by Hellenic Foundation of Scholarships (IKY), through a fellowship awarded to Dr. Antonis D. Lampidonis, M.Sc., Ph.D. References Anthonsen, M.W., Rönnstrand, L., Wernstedt, C., Degerman, E., Holm, C., 1998. Identification of novel phosphorylation sites in hormone-sensitive lipase that are phosphorylated in response to isoproterenol and govern activation properties in vitro. J. Biol. Chem. 273 (1), 215–221. Brasaemle, D.L., Levin, D.M., Adler-Wailes, D.C., Londos, C., 2000. The lipolytic stimulation of 3T3-L1 adipocytes promotes the translocation of hormone-sensitive lipase to the surfaces of lipid storage droplets. Biochim. Biophys. Acta 1483, 251–262. Brenner, S., 1988. The molecular evolution of genes and proteins: a tale of two serines. Nature 334 (6182), 528–530. Contreras, J.A., Holm, C., Martin, A., Gaspar, M.L., Lasuncion, M.A., 1994. Presence of hormone-sensitive lipase mRNA in J774 macrophages. Isr. J. Med. Sci. 30, 778–781. Contreras, J.A., Karlsson, M., Østerlund, T., Laurell, H., Svensson, A., Holm, C., 1996. Hormone-sensitive lipase is structurally related to acetylcholinesterase, bile saltstimulated lipase, and several fungal lipases. J. Biol. Chem. 271 (49), 31426–31430. Cordle, S.R., Colbran, R.J., Yeaman, S.J., 1986. Hormone-sensitive lipase from bovine adipose tissue. Biochim. Biophys. Acta 887 (1), 51–57. De Simone, G., et al., 2001. The crystal structure of a hyper-thermophilic carboxylesterase from the archaeon Archaeoglobus fulgidus. J. Mol. Biol. 314, 507–518. Egan, J.J., Greenberg, A.S., Chang, M.K., Wek, S.A., Moos Jr., M.C., Londos, C., 1992. Mechanism of hormone-stimulated lipolysis in adipocytes: translocation of hormone-sensitive lipase to the lipid storage droplet. Proc. Natl. Acad. Sci. U. S. A. 89, 8537–8541. Fredrikson, G., Strålfors, P., Nilsson, N.Ö., Belfrage, P., 1981. Hormone-sensitive lipase of rat adipose tissue. Purification and some properties. J. Biol. Chem. 256, 6311–6320. Garton, A.J., Campbell, D.G., Cohen, P., Yeaman, S.J., 1988. Primary structure of the site on bovine hormone-sensitive lipase phosphorylated by cyclic AMP-dependent protein kinase. Fed. Eur. Biochem. Soc. 229 (1), 68–72. Garton, A.J., Campbell, D.G., Carling, D., Hardie, D.G., Colbran, R.J., Yeaman, S.J., 1989. Phosphorylation of bovine hormone-sensitive lipase by the cAMP-activated protein kinase. Eur. J. Biochem. 179, 249–254. Gnunmer, R.R., 1991. Effect of feed on the composition of milk fat. J. Dairy Sci. 74, 3244–3257. Greenberg, A.S., et al., 2001. Stimulation of lipolysis and hormone-sensitive lipase via the extracellular signal-regulated kinase pathway. J. Biol. Chem. 276, 45456–45461.

Guex, N., Peitsch, M.C., 1997. SWISS-MODEL and the Swiss-PdbViewer: an environment for comparative protein modeling. Electrophoresis 18, 2714–2723. Haemmerle, G., et al., 2002. Hormone-sensitive lipase deficiency in mice causes diglyceride accumulation in adipose tissue, muscle and testis. J. Biol. Chem. 277 (7), 4806–4815. Harbitz, I., Langset, M., Ege, A.G., Hoyheim, B., Davies, W., 1999. The porcine hormonesensitive lipase gene: sequence, structure, polymorphisms and linkage mapping. Anim. Genet. 30, 10–15. Holm, C., 2003. Molecular mechanisms regulating hormone-sensitive lipase and lipolysis. Biochem. Soc. Trans. 31, 1120–1124. Holm, C., Belfrage, P., Fredrikson, G., 1987. Immunological evidence for the presence of hormone-sensitive lipase in rat tissues other than adipose tissue. Biochem. Biophys. Res. Commun. 148, 99–105. Holm, C., et al., 1988. Hormone-sensitive lipase: sequence, expression, and chromosomal localization to 19 cent-q13.3. Science 241, 1503–1506. Holm, C., Fredrikson, G., Sundler, R., Belfrage, P., 1990. Incorporation of hormonesensitive lipase into phosphatidylcholine vesicles. Lipids 25, 254–259. Holm, C., Davis, R.C., Østerlund, T., Schotz, M.C., Fredrikson, G., 1994. Identification of the active serine of hormone-sensitive lipase by site-directed mutagenesis. FEBS Lett. 344, 234–238. Holm, C., Osterlund, T., Laurell, H., Contreras, J.A., 2000. Molecular mechanisms regulating hormone-sensitive lipase and lipolysis. Annu. Rev. Nutr. 20, 365–393. Holst, L.S., et al., 1996. Molecular cloning, genomic organization and expression of a testicular isoform of hormone-sensitive lipase. Genomics 35, 441–447. Jenkins, C.M., Mancuso, D.J., Yan, W., Sims, H.F., Gibson, B., Gross, R.W., 2004. Identification, cloning, expression, and purification of three novel human calciumindependent phospholipase A2 family members possessing triacylglycerol lipase and acylglycerol transacylase activities. J. Biol. Chem. 279, 48968–48975. Jepson, C.A., Yeaman, S.J., 1993. Expression of hormone-sensitive lipase in macrophage foam cells. Biochem. Soc. Trans. 21, 232 S. Jones, D.T., 1999. Protein secondary structure prediction based on position-specific scoring matrices. J. Mol. Biol. 292, 195–202. Klannemark, M., et al., 1998. The putative role of the hormone-sensitive lipase gene in the pathogenesis of Type II diabetes mellitus and abdominal obesity. Diabetologia 41, 1516–1522. Kozak, M., 1981. Possible role of flanking nucleotides in recognition of the AUG initiator codon by eukaryotic ribosomes. Nucleic Acids Res. 9, 5233–5262. Kraemer, F.B., Shen, W.J., 2002. Hormone-sensitive lipase: control of intracellular tri(di-)acylglycerol and cholesteryl ester hydrolysis. J. Lipid Res. 43, 1585–1594. Laemmli, U.K., 1970. Cleavage of structural proteins during the assembly of the head of the bacteriophage T4. Nature 227, 680–685. Langfort, J., et al., 1998. Hormone-sensitive lipase (HSL) expression and regulation in skeletal muscle. Adv. Exp. Med. Biol. 441, 219–228. Langfort, J., Donsmark, M., Ploug, T., Holm, C., Galbo, H., 2003. Hormone-sensitive lipase in skeletal muscle: regulatory mechanisms. Acta Physiol. Scand. 178, 397–403. Langin, D., Laurell, H., Holst, L.S., Belfrage, P., Holm, C., 1993. Gene organization and primary structure of human hormone-sensitive lipase: possible significance of a sequence homology with a lipase of Moraxella TA144, an Antarctic bacterium. Proc. Natl. Acad. Sci. U. S. A. 90, 4897–4901. Langin, D., et al., 2005. Adipocyte lipases and defect of lipolysis in human obesity. Diabetes 54 (11), 3190–3197. Levitt, R.C., Liu, Z., Nouri, N., Meyers, D.A., Brandriff, B., Mohrenweiser, H.M., 1995. Mapping of the gene for hormone-sensitive lipase (LIPE) to chromosome 19q13.1q13.2. Cytogenet. Cell Genet. 69 (3–4), 211–214. Li, Z., Sumida, M., Birchbauer, A., Schotz, M.C., Reue, K., 1994. Isolation and characterization of the gene for mouse hormone-sensitive lipase. Genomics 24, 259–265. Martín-Hidalgo, A., Huerta, L., Álvarez, N., Alegría, G., del Val Toledo, M., Herrera, E., 2005. Expression, activity and localization of hormone-sensitive lipase in rat mammary gland during pregnancy and lactation. J. Lipid Res. 46, 658–668. Ollis, D.L., et al., 1992. The alpha / beta hydrolase fold. Protein Eng. 5, 197–211. Østerlund, T., et al., 1996. Domain-structure analysis of recombinant rat hormonesensitive lipase. Biochem. J. 319, 411–420. Osuga, J., et al., 1997. Suppression of neutral cholesterol ester hydrolase activity by antisense DNA of hormone-sensitive lipase. Biochem. Biophys. Res. Commun. 233, 655–657. Palmquist, D.L., 1984. Use of fats in diets for lactating cows. In: Wiseman, J. (Ed.), Fats in Animal Nutrition. Buttenvolths, London, England, p. 357. Sambrook, J., Fritsch, E.F., Maniatis, T., 1989. Molecular Cloning: A Laboratory Manual, 2nd Ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY, USA. Shen, W.-J., Patel, S., Natu, V., Kraemer, F.B., 1998. Mutational analysis of structural features of rat hormone-sensitive lipase. Biochemistry 37, 8973–8979. Shen, W.-J., Sridhar, K., Bernlohr, D.A., Kraemer, F.B., 1999. Interaction of rat hormonesensitive lipase with adipocyte lipid-binding protein. Proc. Natl. Acad. Sci. U. S. A. 96, 5528–5532. Small, C.A., Garton, A.J., Yeaman, S.J., 1989. The presence and role of hormone-sensitive lipase in heart muscle. Biochem. J. 258, 67–72. Smith, M.G., Garton, A.J., Aitken, A., Yeaman, S.J., 1996. Evidence for a multi-domain structure for hormone-sensitive lipase. FEBS Lett. 396, 90–94. Stralfors, P., Olsson, H., Belfrage, P., 1987. The Enzymes, vol. 18. Academic Press, New York, pp. 147–177. Su, C.-L., Sztalryd, C., Contreras, J.A., Holm, C., Kimmel, A.R., Londos, C., 2003. Mutational analysis of the hormone-sensitive lipase translocation reaction in adipocytes. J. Biol. Chem. 278 (44), 43615–43619. Syu, L.J., Saltiel, A.R., 1999. Lipotransin: a novel docking protein for hormone-sensitive lipase. Mol. Cell 4, 109–115.

A.D. Lampidonis et al. / Gene 416 (2008) 30–43 Sztrolovics, R., Wang, S.P., Lapierre, P., Chen, H.S., Robert, M.-F., Mitchell, G.A., 1997. Hormone-sensitive lipase (Lipe): sequence analysis of the 129Sv mouse Lipe gene. Mamm. Genome 8, 86–89. Tsujita, Τ., Ninomiya, H., Okuda, H., 1989. p-nitrophenyl butyrate hydrolyzing activity of hormone-sensitive lipase from bovine adipose tissue. J. Lipid Res. 30 (7), 997–1004. Villena, J.A., Roy, S., Sarkadi-Nagy, E., Kim, K.-H., Sul, H.S., 2004. Desnutrin, an adipocyte gene encoding a novel patatin domain-containing protein, is induced by fasting and glucocorticoids: ectopic expression of desnutrin increases triglyceride hydrolysis. J. Biol. Chem. 279, 47066–47075.

43

Warden, C.H., et al., 1993. Chromosomal localization of lipolytic enzymes in the mouse: pancreatic lipase, colipase, hormone-sensitive lipase, hepatic lipase and carboxyl ester lipase. J. Lipid Res. 34, 1451–1455. Wei, Y., et al., 1999. Crystal structure of brefeldin A esterase, a bacterial homolog of the mammalian hormone-sensitive lipase. Nat. Struct. Biol. 6 (4), 340–345. Zimmermann, R., et al., 2004. Fat mobilization in adipose tissue is promoted by adipose triglyceride lipase. Science 306, 1383–1386.