extfieldsofstudy: Biology Medicine

s2fieldsofstudy: Biology Medicine

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High-risk HPV E5-induced cell fusion: a critical initiating event in the early stage of HPV-associated cervical cancer

Background Cervical cancer is strongly associated with high-risk human papillomavirus (HPV) and viral oncoproteins E5, E6 and E7 can transform cells by various mechanisms. It is proposed that oncogenic virus-induced cell fusion may contribute to oncogenesis if p53 or apoptosis is perturbed simultaneously. Recently, HPV-16 E5 was found to be necessary and sufficient for the formation of tetraploid cells, which are frequently found in precancerous cervical lesions and its formation is strongly associated with HPV state. Presentation of the hypothesis We propose that high-risk HPV E5-induced cell fusion is a critical initiating event in the early stage of HPV-associated cervical cancer. Testing the hypothesis Our hypothesis can be tested by comparing the likelihood for colony formation or tumorigenic ability in nude mice between normal HaCaT cells expressing all three oncogenic proteins and E5-induced bi-nucleated HaCaT cells expressing E6 and E7. Moreover, investigating premature chromosome condensation (PCC) in HPV-positive and negative precancerous cervical cells is another way to assess this hypothesis. Implication of the hypothesis This viewpoint would change our understanding of the mechanisms by which HPV induces cervical cancer. According to this hypothesis, blocking E5-induced cell fusion is a promising way to prevent the progression of cervical cancer. Additionally, establishment of a role of cell fusion in cervical carcinogenesis is of reference value for understanding the pathogenesis of other virus-associated cancers.

Background
Cervical cancer progression is strongly associated with infection of high-risk human papillomavirus (HPV) (e.g., , which are detected in nearly all cervical cancers [1]. HPV is a small, nonenveloped DNA virus expressing three key oncoproteins: E5, E6 and E7, which possess the ability of transforming certain human cells in vitro and are considered to be associated with cervical carcinogenesis in vivo [2][3][4][5]. E6 and E7 are well known for their ability to inhibit the function of tumor suppressors p53 and pRb, respectively [6]. E5 has weak oncogenic properties which occur through increasing epidermal growth factor receptor (EGFR) and inhibiting the expression of major histocompatibility complex (MHC)-I and MHC-II on the plasma membrane [7]. Coexpression of E5 with either E6 or E7, however, promotes transformation by either oncoprotein alone [8].
Recently, the view that oncogenic virus-induced cell fusion may contribute to oncogenesis is appealing as all well-known human oncogenic viruses, including HPV, Hepatitis B virus, Hepatitis C virus, Epstein-Barr virus, Kaposi sarcoma virus and human T-lymphotropic virus type 1, have fusogenic activity [9][10][11]. Researches show that although most tetraploid cells resulting from nononcogenic-induced cell fusion would undergo p53dependent cell cycle arrest or apoptosis, they, however, survive and might be more prone to chromosomal instability (CIN) if p53 or apoptosis is perturbed [12,13]. It is notable that all oncogenic viruses mentioned above also possess proteins with these abilities [9]. As to HPVassociated cervical cancer, this mechanism may be operative as HPV-16 E5 was recently found to be necessary and sufficient for the formation of tetraploid cells [10,11], which are frequently found in precancerous cervical lesions and its formation is strongly associated with HPV state [14,15].

Tetraploid cells in precancerous cervical lesions
Tetraploid cervical cells are often observed in precancerous lesions, and it has been established as a prognostic factor that allows to estimate the relative progression risk into more advanced lesions [15]. Moreover, aneuploid cells are frequently observed in these precancerous lesions. It is, therefore, proposed that a sequential pattern of chromosomal aberrations occurs during cervical carcinogenesis, where aneuploidy develops through chromosomal loss from a tetraploid intermediate.

Another study investigated both tetraploid cervical cells and HPV state in precancerous cervical cells, results
showed that tetraploid cervical cells were elevated in women diagnosed as either atypical squamous cells of undetermined significance (ASCUS)/HPV-positive or low-grade squamous intraepithelial lesion (LSIL)/HPVpositive as compared with normal/HPV-negative women, indicating that formation of tetraploid cells is obviously associated with HPV infection [14].

HPV-16 E5 induces cell-cell fusion
HPV-16 E5 has all the characteristics of fusogenic proteins, including localization to the plasma membrane, high level of hydrophobicity, and the ability for dimmers [3]. Until recently, HPV-16 E5, however, has been identified to be necessary and sufficient to induce cell-cell fusion [10]. It is worth mentioning that HPV-16 E5 must be expressed on both cells for cell fusion to occur [11]. It is also found that tetraploid cells were produced with greater than a 3-fold frequency upon introduction of the HPV-16 genome into spontaneously immortalized human keratinocytes (HaCaT) as compared to cells transfected with an HPV-16 genome harboring a mutant E5 gene. By contrast, low-risk HPV-6b E5 could not induce cell fusion [10].
Based on the observations mentioned above, we hypothesize that high-risk HPV E5-induced cell fusion is a critical event in the early stage of HPV-associated cervical cancer.

Presentation of the hypothesis
The fact that aneuploid cells are frequently observed in precancerous lesions with elevated proportion of tetraploid cells, the formation of which is obviously associated with HPV infection [14] suggests that formation of tetraploid cells is a critical event in cervical carcinogenesis, but the detail formation mechanism of tetraploidy is not clear. It is reported that expression of either HPV E6 or E7 alone is sufficient to deregulate cytokinesis and consequently produce tetraploid cells [16,17]. However, Hu et al. demonstrated that the formation of these cells is primarily attributed to E5 and E5-induced cell fusion, rather than E6, E7 and cytokinesis failure [10].
Tetraploid cells formed by accident can not undergo normal mitosis which would trigger p53-dependent cell cycle arrest or apoptosis [12,13], whereas, oncogenic virus-induced cell fusion is sufficient to induce CIN when fusion occurs concomitantly with expression of viral oncoproteins capable of perturbing p53 or apoptosis [12]. Consistently, although most tetraploid cells die out, whereas, coexpression of HPV-16 E6/E7 enhances the proliferation of these cells and the likelihood for colony formation elevates 3-fold [10].
Based on the observations mentioned above, we propose that high-risk HPV E5-induced cell fusion may play a critical role in the early stage of HPV-associated cervical cancer. However, it is widely accepted that increasingly deregulated expression of the E6-E7 oncogenes of high-risk HPVs has been identified as the major transforming factor in the pathogenesis of cervical dysplasia and derived cancers [2]. In fact, these two mechanisms are not mutually exclusive as E5 functions only in the early stage, whereas, E6 and E7 act throughout the carcinogenesis. Additionally, it is notable that expression levels of all three oncoproteins in host cell are low for tight restriction in the early stage, and expression of E5 is nearly not detectable in all late stages for integration [3]. Moreover, the ability of E6 and E7 to transform various cells have been demonstrated only in overexpression systems, therefore, whether it also occurs in natural settings is not known. We also notice that in vitro [18,19] and clinic studies [20] reveal that chromosomal instability and aneuploidization seem to precede and favor integration of HPV genomes, which in turn leads to expression of viral-cellular fusion transcripts and further enhances expression of the E6-E7 genes, which renders transformed cells strong growth advantages [21].

Testing the hypothesis
In order to determine whether high-risk HPV E5induced cell fusion is important for initial transformation, we can compare the likelihood for colony formation or tumorigenic ability in nude mice between normal HaCaT cells expressing all three oncogenic proteins and E5-induced bi-nucleated HaCaT cells expressing E6 and E7. In our hypothesis, cell fusion and cell cycle deregulation are two key events for initiation of transformation. Therefore, inhibition of cell fusion should significantly decrease the likelihood for transformation. Moreover, investigating premature chromosome condensation (PCC) in HPV-positive and negative precancerous cells is another way to test this hypothesis. PCC refers to condensation of interphase chromosomes following fusion between an interphase and a mitotic cell [22], and it can be used as a tool to detect the existence of cell fusion [23].

Implication of the hypothesis
Understanding the cause of cancer is critical for effective diagnosis, prevention and therapy. The recent view that human oncogenic virus-induced cell fusion can initiate cancer is an appealing mechanism. In this article, we propose that high-risk HPV E5-induced cell fusion is an important event in the early stage of HPV-associated cervical cancer. This viewpoint will change our understanding of the mechanisms by which HPV induces cervical cancer. According to this hypothesis, blocking HPV E5-induced cell fusion is a promising way to prevent the progression of cervical cancer. Additionally, establishment of a role of cell fusion in carcinogenesis of cervical cancer is of reference value for understanding the pathogenesis of other virus-associated cancers.


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Synthetic biology approaches in drug discovery and pharmaceutical biotechnology

Synthetic biology is the attempt to apply the concepts of engineering to biological systems with the aim to create organisms with new emergent properties. These organisms might have desirable novel biosynthetic capabilities, act as biosensors or help us to understand the intricacies of living systems. This approach has the potential to assist the discovery and production of pharmaceutical compounds at various stages. New sources of bioactive compounds can be created in the form of genetically encoded small molecule libraries. The recombination of individual parts has been employed to design proteins that act as biosensors, which could be used to identify and quantify molecules of interest. New biosynthetic pathways may be designed by stitching together enzymes with desired activities, and genetic code expansion can be used to introduce new functionalities into peptides and proteins to increase their chemical scope and biological stability. This review aims to give an insight into recently developed individual components and modules that might serve as parts in a synthetic biology approach to pharmaceutical biotechnology.

Introduction
Nature provides a reservoir of active compounds that help us to fight and cure disease (Fig. 1). We have learned to exploit this reservoir by identifying and isolating the active compounds, by developing strategies for their chemical synthesis and by improving their biological activity. However, this process is laborious and very expensive. The largescale isolation of drugs is often hindered by the low amount of material produced by the natural source, and the complexity of the molecules makes their total chemical synthesis costly to unprofitable. Transferring the ability to biosynthesize these compounds from the source organisms to genetically friendlier hosts would greatly facilitate their production. Genome sequencing efforts now provide the genetic information of entire metabolic pathways necessary to recombinantly produce pharmaceutically relevant molecules. However, biosynthetic pathways are often highly intertwined, regulated by extensive feedback mechanisms and require the correct organization of the individual entities in space and time. The simple transfer of genetic information from one organism to another is therefore rarely sufficient to confer the biosynthetic capability. Thus, we need to understand the properties of individual components as well as the properties of entire systems and eventually learn to recombine artificial biosynthetic networks in new hosts in a predictable, efficient manner. The design of life with novel properties from individual modules in a way resembling the creation of new machines in modern engineering is the ultimate aim of the emergent discipline of synthetic biology.
In its most stringent definition, synthetic biology is the application of the principles of engineering to the construction of life with desired properties in a rational and systematic way (Serrano 2007). Synthetic biology therefore uses well-characterized, modular parts that can be put together to create new functionality in a directed, predictable manner (Kiel et al. 2010;Ninfa 2010;Topp and Gallivan 2010). Several recent advances in DNA technology encourage the followers of this new discipline: First, inexpensive DNA synthesis enables the facile production of codon-usage optimized genes and other genetic elements, such as promoters and regulatory sequences. This technology has recently been improved towards the synthesis of an entire genome, which was subsequently used to substitute the genome of a living cell Gibson et al. 2008a, b;Lartigue et al. 2007Lartigue et al. , 2009. Second, rapid, next-generation DNA sequencing technologies provide vast amounts of genetic information. Entire genome sequences of many organisms are now publicly available and can be mined electronically for relevant genes (Benson et al. 2009). Third, wellcharacterized genetic modules are being developed, which can be combined rationally to create a new function (Lu et al. 2009). The Registry of Standard Biological Parts (http://parts. mit.edu) is a continuously growing collection of standardized parts ("BioBricks") that are being used to rationally create organisms with desired properties (Voigt 2006). Directed evolution has become a widespread strategy to adapt existing parts to novel functions. For example, the directed evolution of tRNAs and amino acyl-tRNA synthetases (aaRS) to decode novel, unnatural amino acids has been used to expand the genetic code of several organisms (Chin et al. 2003;Liu et al. 2007;Wang et al. 2001) and allows genetically programming the incorporation of unprecedented functionality into proteins in vivo (Xie and Schultz 2006).
With increasing experience in the reprogramming of microorganisms, entirely new types of therapy might become feasible in the future. Bacteria might no longer be limited to the mere production of drugs but will also be able to deliver them to the site of action. In this review, we want to give an overview of recently developed individual components and of the attempts to recombine them for the production of modules of biotechnological and pharmaceutical interest. We are trying to give an impression of how these individual modules might eventually be used as parts in a synthetic biology approach to pharmaceutical biotechnology. Producing compound libraries directly in vivo would have several advantages. First, maintaining and amplifying such a library is simple and can be done by cultivating cells. Second, the screening process is facilitated because the genetic information remains directly coupled to the compound. Hence, the selected compound can be identified by analysing the genetic information of the corresponding cell. Third, an intracellular genetic selection can directly assay for effects on enzymatic activity, bypassing inherent limitations of in vitro assays. Fourth, the selection is performed in the context of a living cell requiring an enhanced level of selectivity of the drug for its target. Finally, potential problems with solubility and uptake of the compound are circumvented.

Cyclic peptide libraries
Many important natural compounds are modified cyclic peptides, for example, antibiotics like vancomycin (Fig. 1a), the immunosuppressant drug cyclosporin (Fig. 1b) and fungal toxins phalloidin or actinomycin D (Katsara et al. 2006). Cyclization renders peptides resistant to cellular degradation and restricts conformational freedom, potentially improving binding affinity and specificity. Using a split intein approach, the Benkovic lab has developed a method to produce libraries of cyclic peptides in Escherichia coli (Scott et al. 1999). A number of promising candidates have since been identified in libraries generated this way (Horswill et al. 2004;Tavassoli and Benkovic 2005;Tavassoli et al. 2008). Natural cyclic peptides contain many modified, non-coding amino acids. This increases the diversity of these compounds and is crucial for their biological activity. As detailed below, the artificial expansion of the genetic code allows the cotranslational incorporation of unnatural amino acids (Wang et al. 2001;Xie and Schultz 2006). Combining the Benkovic approach to the production of cyclic peptides with the incorporation of unnatural amino acids would vastly increase the diversity of cyclic peptides that can be produced in vivo.

Polyketide synthases-natural synthetic biology
Another important class of natural products are the polyketides. Polyketides are synthesized by large multienzyme complexes, the polyketide synthases. These "as-sembly lines" are built from modular components that catalyze the formation of the carbon chain of the final product in a stepwise manner (Fig. 2). The modular composition of polyketide synthases makes them an ideal playground for synthetic biologists. Their individual modules can be split and recombined to form active hybrid enzymes (Watanabe et al. 2003b). The combinatorial recombination of individual modules has been achieved, and the new enzymes were shown to successfully catalyze the formation of polyketides in E. coli (Menzella et al. 2005). This approach can potentially be used to produce libraries of polyketides with novel biological activities in vivo. Presently, simple recombination of different modules often produces inactive synthases because the transfer of intermediates between modules might be blocked or the connectivity between modules disturbed. Growing information on the structure of individual modules and their connectivity (Alekseyev et al. 2007;Keatinge-Clay and Stroud 2006;Tang et al. 2006) together with increasing experimental experience will help to develop predictive algorithms to rationally design synthases for "unnatural" polyketides in a combinatorial biosynthetical approach (Khosla et al. 2009). If a system to produce large libraries of hybrid enzymes is connected to a selectable output (Yin et al. 2007), active clones can be identified, even if their frequency in the library is very low (Menzella and Reeves 2007). A combination of computer-assisted prediction, combinatorial library design guided by structural information and selection might eventually develop into the mainstream of drug discovery.

De novo creation of metabolic pathways
The de novo design of biosynthetic pathways to expand the biosynthetic potential of microorganisms is an intriguing route towards compounds for which natural pathways have not been elucidated or which are not of natural origin (Prather and Martin 2008). A "retro-biosynthetic" approach, analogous to retrosynthesis in organic chemistry, uses enzymes with evolved substrate specificity to create artificial biosynthetic pathways de novo. Many natural enzymes show high selectivity for their substrates, which is the result of divergent evolution from promiscuous precursor proteins (O'Brien and Herschlag 1999). However, there are examples of natural enzymes with broad substrate specificity. Starting from such a promiscuous enzyme, the sesquiterpene synthase γ-humulene synthase, Keasling and co-workers were able to recapitulate this evolutionary process and create seven specific and active enzymes that use different reaction pathways and produce different products (Yoshikuni et al. 2006). These enzymes could be used in the future to design biosynthetic pathways for unnatural terpenoids.
Similarly, prenylation is an important modification occurring on natural products, like naphterpin, conferring anti-cancer, anti-viral or anti-microbial activity to the molecules (Botta et al. 2005). Prenyltransferases, the enzymes responsible for the derivatization, have been identified and characterized (Kuzuyama et al. 2005). These enzymes have a broad substrate spectrum and might form a starting point for the evolution of enzymes for regiospecific prenylation of aromatic small molecules. Engineered enzymes might eventually allow us to create enzymatic pathways de novo and to produce small molecule libraries of differently modified compounds of the same scaffold structure (Dietrich et al. 2009;Yoshikuni et al. 2006).

Expanding the chemistry of life
The cellular environment imposes a constraint to the chemical scope of reactions that can be used to create in vivo compound libraries. Many reactions familiar to synthetic organic chemists require elevated temperature or are incompatible with aqueous environments. This limitation may be overcome in the future by the computer-assisted, directed evolution of enzymes (Kaplan and DeGrado 2004) Fig. 2 Microbial polyketide synthases are modular assembly lines that fit polyketides together from monomeric building blocks. In the first step of the reaction, the starter module is acylated with the first unit (1). This unit is subsequently transferred to the first of a row of extender modules that catalyze the elongation of the polyketide. All extender modules contain at least a ketosynthase (KS), an acyl transferase (AT) and an acyl carrier protein (ACP) domain. In each step, the KS receives the acyl group unit from the ACP of the preceding module and the AT adds an appropriate extender unit from its CoA ester to the prosthetic group of the ACP (2). The KS then catalyzes a decarboxylative Claisen condensation between the acylated KS and the extender unit to give a β-keto-acylated acyl carrier protein (3). Other modules may be included to further modify the growing polyketide chain. Upon completion of the elongation process, the polyketide chain is cleaved from the polyketide synthase by the thioesterase (TE) of the termination module (4) reactions. Using several enzymes as scaffolds, Baker and coworkers (Rothlisberger et al. 2008) employed computer-aided design to engineer enzymes able to catalyze a Kemp elimination reaction for which no natural enzyme is yet known. It is thus conceivable that the limits to the chemistries available in vivo will be shifted in the future, thereby expanding the chemical scope of in vivo compound libraries.
The diversity in the chemical nature of proteins and peptides has been tremendously increased by the invention of strategies to introduce unnatural amino acids in vivo. Peter Schultz and his co-workers have artificially expanded the genetic code of several organisms (Chin et al. 2003;Liu et al. 2007;Wang et al. 2001) and succeeded in incorporating a wide range of unnatural amino acids (Xie and Schultz 2006). The basic principle of genetic code expansion is the expression of an additional pair of tRNA and its cognate aaRS in the host system (Fig. 3). Both, tRNA and aaRS, must not be substrates for the host aaRSs and tRNAs to ensure correct translation of host proteins. Under these prerequisites, the tRNA can be mutated to decode amber stop codons, for example, and the aaRS can be evolved to specifically recognize unnatural amino acids. This system allows the incorporation of additional, unnatural amino acids in response to amber stop codons. In the past decade, the list of unnatural amino acids that can be incorporated in this way has grown continuously (Neumann et al. 2008;Nguyen et al. 2009b;Xie and Schultz 2006). Recent developments of amber and quadruplet suppressor ribosomes now allow the efficient incorporation of multiple different unnatural amino acids into the same protein (Neumann et al. 2010;Wang et al. 2007). These unnatural amino acids have side chains with unusual chemical properties and reactivities and could be used to increase the chemical diversity of peptides and proteins.

Drug screening
The quality of an assay system has an enormous impact on the outcome of a screen. In order to fully profit from the advantages of in vivo compound libraries, it is necessary to develop intracellular assays, like a colour signal or a genetically selectable phenotype, to detect or select positive clones. The continuously increasing number of biosensors could serve as such detector modules.
The Benkovic group has developed a method for identifying small molecule inhibitors of protein-protein interactions by combining a reverse two-hybrid system with the split intein-mediated formation of cyclic peptide libraries (Horswill et al. 2004). Using this system, cyclic peptides were identified that modulate the interaction between the two subunits of ribonucleotide reductase (Horswill et al. 2004), HIV-Gag and TSG101 (Fig. 1c) (Tavassoli et al. 2008) and AICAR transformylase homodimerization (Tavassoli and Benkovic 2005). A similar system that combines in vivo cyclic peptide libraries with a Fig. 3 The principle of genetic code expansion. The basic principle of genetic code expansion is the heterologous expression of an additional tRNA/aminoacyl-tRNA synthetase (aaRS) pair in the host cell. This pair is orthogonal to the host's translational machinery, meaning that the tRNA is not a substrate for endogenous synthetases and that the aaRS does not charge host tRNAs. The orthogonal tRNA is mutated to decode amber stop codons and the orthogonal aaRS is evolved to specifically recognize an unnatural amino acid (UAA) supplied with the growth medium. Cells with such an expanded genetic code are able to produce proteins with the UAA incorporated at a position encoded by an amber codon in the mRNA fluorescent reporter assay was developed by the same group and used to identify antibacterial cyclic peptide inhibitors of ClpXP protease (Cheng et al. 2007).
Tsien and co-workers created the Ca 2+ -sensor "Cameleon" (Miyawaki et al. 1997) by fusing calmodulin to a calmodulinbinding peptide and flanking the construct with two fluorescent proteins with overlapping spectral properties. Ca 2+ -dependent binding of calmodulin to its own peptide results in a predictable conformational rearrangement that leads to a change in energy transfer between the two fluorescent proteins. Biosensors of this sort are being used extensively to monitor the intracellular concentration and dynamics of ions and small molecules and the activation state of signalling proteins (Balla 2009). Similar sensors could be designed for metabolites of interest and combined with systems to produce small molecule libraries in vivo in order to identify strains and conditions that allow production of pharmacologically relevant compounds.
Optimization of engineered metabolic pathways depends on our ability to measure the production of intermediates efficiently. Mass spectrometry can be used to quantify the relevant metabolites, but faster real-time assays allow the identification of optimal growth condition or the screening of mutant libraries in high-throughput formats. Mevalonate, for example, is an important precursor in the biosynthesis of isoprenoids. Keasling and co-workers (Pfleger et al. 2007) engineered an auxotrophic strain of E. coli into a mevalonate biosensor by expressing GFP. The strain reports on the mevalonate concentration in the growth medium through a change in growth rate. This biosensor strain can now be used to measure mevalonate in high-throughput formats.
In a more direct approach, reporter proteins could be created by redesigning the affinities of binding proteins, like maltose binding protein (MBP), for ligands of interest. MBP is a member of the periplasmic binding protein superfamily that mediates chemosensory processes in a wide range of bacteria. This protein family has evolved to bind a variety of ligands, such as amino acids, carbohydrates and ions, naturally and has served as starting point for the directed evolution of biosensors, receptors and enzymes (Dwyer and Hellinga 2004). Upon ligand binding, these proteins undergo a significant conformational change (induced fit), which has been exploited to monitor the event by positioning environmentally sensitive fluorophores within the binding site (endosteric placement), at the rim of the binding site (peristeric placement) or in a crevice that opens and closes with ligand binding (allosteric placement) (de Lorimier et al. 2002). Alternatively, GFP variants have been conjugated to the N-and C-termini of periplasmic-binding proteins in order to monitor ligand binding by fluorescence energy transfer (Fehr et al. 2002;Fehr et al. 2003;Schultz and Moini 2003). Computational design has been successfully employed to create biosensor proteins with novel affinities for small molecules, such as trinitrotoluene, L-lactate and serotonin (Looger et al. 2003). These engineered sensors were incorporated into synthetic bacterial signal transduction pathways to regulate gene expression in response to extracellular stimuli (Looger et al. 2003).
G-protein coupled receptors (GPCRs) are the target of the majority of pharmaceuticals. This membrane protein family is stimulated by extracellular ligands and initiates a signalling cascade on the cytosolic side of the membrane upon agonist binding. The study of GPCRs in their natural environment is usually complicated by the presence of multiple different receptors on a single cell combined with the tremendous efforts necessary for its genetic manipulation. Pausch and colleagues transplanted a mammalian GPCR into an engineered Saccharomyces cerevisiae strain and connected it to a downstream cascade controlling the expression of the his3 gene (Price et al. 1995). Activation of the receptor permitted the cells to grow in the absence of histidine. This approach has since been used extensively to study GPCR function and to identify ligands and antagonists (Brown et al. 2000;Minic et al. 2005;Sachpatzidis et al. 2003).
These examples demonstrate that it is possible to create modules for the detection of a molecule of interest by rational design. A synthetic biology approach to the production of pharmaceuticals should employ these sensors to control the expression level of individual parts or connect them to a selectable output. In a case study, Farmer and Liao coupled the E. coli Ntr regulon, which senses the intracellular metabolite acetylphosphate, to the expression of key enzymes in the pathway for the recombinant biosynthesis of lycopene (Farmer and Liao 2000). This approach significantly enhanced lycopene production and simultaneously reduced the toxic side effects caused by metabolic imbalance. Riboswitches, RNA elements with the ability to control the translation of a coupled open reading frame upon binding specifically to certain ligands, are another class of parts that can be used to control metabolism (Famulok et al. 2007). Even before their discovery in nature, artificial riboswitches had been designed to control the expression of reporter genes in response to the binding of small molecules, such as Hoechst dyes or tetracycline Suess et al. 2004;Suess et al. 2003;Werstuck and Green 1998). These elements could be used to couple gene expression on the translational level to the metabolic state of the cell. Similar strategies might form the basis of highthroughput screening systems to identify engineered strains that are able to biosynthesize the molecule of interest.

Biosynthesis of pharmaceuticals
Recombinant DNA technology has been used for more than three decades to engineer bacteria with the ability to produce pharmacologically important molecules like insulin (Goeddel et al. 1979). Progress has been made in recent years towards the heterologous production of important natural products, like terpenoids, polyketides, nonribosomal peptides and alkaloids. Synthetic biology approaches are being employed to fine-tune the levels and activities of individual steps and components. Here, we present examples of recent successes of this strategy.

Terpenoids
Terpenoids are a large and diverse class of natural products with important applications in human health. Well-known examples are the anti-malarial drug artemisinin (Fig. 1d) (Tan et al. 1999), the anti-cancer agent paclitaxel (taxol) (Jennewein and Croteau 2001) and eleutherobin (Long et al. 1998). A major step forward towards the recombinant production of terpenoids in E. coli was the creation of a platform strain that heterologously expressed the mevalonate pathway of S. cerevisiae (Martin et al. 2003). The metabolic precursors produced by this strain can be further converted into relevant terpenoids by co-expression of the corresponding terpene synthases and modifying enzymes. Building on this work, the Keasling lab has engineered E. coli to produce artemisinic acid, which can be converted chemically into artemisinin (Roth and Acton 1989), with yields of more than 300 mg/l (Chang et al. 2007;Kizer et al. 2008;Newman et al. 2006). Similarly, the same group has recently reprogrammed the metabolism of S. cerevisiae to produce artemisinic acid with titres exceeding 100 mg/l (Ro et al. 2006).
Taxol (paclitaxel) is a well-established anti-neoplastic drug active against a variety of different cancers (Goldspiel 1997). Although the total synthesis of Taxol has been achieved (Kingston et al. 2002), the approach is too expensive for it being commercially practical (Borman 1994). The drug is produced by yew (Taxus) species in about 19 individual enzymatic steps from the isoprenoid precursor geranylgeranyl diphosphate (Jennewein and Croteau 2001;Walker and Croteau 2001). Several Taxol biosynthetic genes have been expressed functionally in S. cerevisiae and used to establish the five-step biosynthesis of an intermediate of Taxol (Dejong et al. 2006;Engels et al. 2008). Efforts are also being undertaken to establish Taxol biosynthesis in E. coli (Huang et al. 2001). However, there are still a number of problems to overcome, like the lack of a complete list of enzymes involved in the production of many important terpenoids, in order to achieve their heterologous total biosynthesis. Still, the achievements are promising and there is hope for cost-efficient production of some very important natural products in the near future.

Polyketides
Polyketides are a large class of important natural products, e.g. erythromycin, epothilone (Fig. 1e) and FK-506. They are produced by microorganisms, most prominently actinomycetes, soil-dwelling bacteria. A significant achievement in the recombinant production of polyketides in E. coli (that naturally does not produce any) was the engineering of the pathway that encodes the production of 6-deoxyerythronolide B (6dEB) (Pfeifer et al. 2001). This strain was created by optimizing precursor production, enzyme engineering and destroying catabolic pathways. Subsequent optimization of the strain has improved the level of 6dEB production to levels obtained from optimized S. coelicolor strains (Pfeifer et al. 2002). Building on this work, heterologous production of several other important polyketides or their precursors has been achieved recently, namely, the anti-cancer drugs epothilone C and D (Mutka et al. 2006) and ansamycin precursors (Rude and Khosla 2006;Watanabe et al. 2003a), aklanoic acid (precursor to several antitumor polyketides, e.g. doxorubicin and aclacinomycin A) (Lee et al. 2005) and aromatic bacterial polyketides (Zhang et al. 2008).

Metabolic fine-tuning
These promising examples illustrate the potential of a synthetic biology approach to the recombinant production of pharmaceutics. There are, however, significant challenges ahead. In many cases, the relevant precursor metabolites are inefficiently produced by the host or bottlenecks are formed during the synthesis of the product when an intermediate is unstable or an enzyme of low activity in the foreign environment. Synthetic biology has the aim to create autonomous circuits for the production of key metabolites allowing the modular combination of part reactions into new pathways. The carefully adjusted expression level of individual components of the pathway is important to optimize flux and to avoid the accumulation of toxic intermediates. Synthetic promoter libraries and a range of engineered constitutive promoters of varying strengths have been created (Alper et al. 2005;Hammer et al. 2006) and could be used to fine-tune the expression levels of all enzymes in a pathway in order to optimize metabolic flux.
The Church lab has recently published a novel technology for multiplex automated genome engineering ("MAGE") to optimize metabolic pathways in E. coli. By feeding cells with synthetic oligonucleotides, which are probably used as Okazaki fragments during replication, the authors were able to mutate and fine-tune ribosomal binding sites genome-wide (Wang et al. 2009). They have used this method to increase the production of lycopene in an engineered E. coli strain by a factor of five within only 3 days of evolution. The same approach might allow optimizing the production of other metabolites if a simple screening assay is available. The use of an automated setup greatly accelerated and increased the throughput of this technology. The development of similar automated systems will have a major impact on the de novo creation of biosynthetic pathways and regulatory networks in the future.
An important reoccurring theme of cellular metabolism is compartmentalization and channelling of metabolites (Ovadi and Srere 2000). This avoids side reactions (especially hydrolysis) of activated intermediates and prevents toxic side effects. A strategy that mimics this organization of enzymes would facilitate the design of efficient metabolic pathways. A first step towards this aim has recently been taken: By fusing the components of the metabolic pathway for mevalonate to peptide motifs, Keasling and co-workers (Dueber et al. 2009) were able to recruit them into synthetic protein scaffolds and to optimize their stoichiometry within the synthetic complex. The activity of the pathway was thereby increased by a factor of 77. Compartmentalization (for example, in protein-based organelles that exist in many bacteria (Yeates et al. 2008)) can also help to establish new chemical reactions by shielding unstable intermediates from a hostile environment or by creating a special phase, e.g. with low or high pH or hydrophobicity.

Bacteria as drug delivery units
In recent years, we have witnessed an explosion of new technologies to manipulate cellular genetic information. Our engineering abilities allow us to design cells with artificial genomes Gibson et al. 2008a, b;Lartigue et al. 2007Lartigue et al. , 2009, promising the advent of customized drug-producing or therapeutic bacteria. It might become possible to equip "synthetic" organisms with just the genetic information necessary for function, while omitting any interfering and potentially dangerous genetic and metabolic material. These cells could be rationally equipped with modular genetic elements to create an organism with a desired phenotype. In a futuristic scenario, genetic devices might be combined to program organisms like robots. First attempts produced newly combined biosensor modules under newly linked genetic circuits (Kobayashi et al. 2004;Looger et al. 2003;Lu et al. 2009;You et al. 2004). For example, a chimeric sensor domain of a phytochrome from cyanobacteria fused to a signal transduction domain of E. coli was used to design an E. coli strain responding to light (Levskaya et al. 2005). Engineering new bacterial sensors to regulate genes in response to a new environmental context can be extended to create other sensors and also reprogram downstream signalling at wish. Promoters, for example, could be built to integrate (multiple) specific signals (Anderson et al. 2007).
This approach can be extended to design live bacteria as targeted delivery systems for live vaccination, as probiotics and anti-tumour agents (Garmory et al. 2003;Martin et al. 2003;Nguyen et al. 2010;Pawelek et al. 2003). Various bacteria invade tumours and have been engineered to destruct them by releasing a chemotherapeutic prodrug (Nemunaitis et al. 2003), via secretion of TNFα (Dang et al. 2001;Nuyts et al. 2001) and cytokines (Murray et al. 1996). However, a synthetic biological approach would allow designing a bacterium rationally with desired capabilities and safety requirements. The Voigt lab constructed an E. coli strain that senses a low-oxygen microenvironment such as found in tumour tissues. Hypoxia is the cue in this particular E. coli strain to upregulate a Yersinia pseudotuberculosis adhesin protein (invasin) sufficient for E. coli to invade mammalian cells. Because invasion is only efficient from a certain cell density, the Voigt lab further equipped this E. coli with a second sensor including another genetic circuit that senses cell density (quorumsensing circuit from Vibrio fischeri) (Anderson et al. 2006). Thus, this E. coli strain should specifically invade tumour cells in a population density-dependent manner.
The type III secretion apparatus, usually used by certain bacteria to pump effector proteins into the eukaryotic host cytoplasm for manipulation, was exploited in a synthetic biology approach to pump spider silk proteins into the medium (Widmaier et al. 2009). Combined with the previous approaches, a (synthetic) bacterial strain equipped with rationally linked sensors and genetic circuits might be able to sense the environment of damaged tissues, specifically to adhere and release therapeutic proteins into the infected cell and thus could kill (or heal) damaged or infected tissues. Refining such approaches together with biological "safety tasks", which can also be synthetic, would be a further step towards the use of safe therapeutic bacteria.

Optimizing drugs
Often the identification of a new drug candidate is only the beginning of a laborious and time-consuming effort to optimize and mature its pharmacokinetic properties.
Small therapeutic proteins, for example, have a short residence time in the bloodstream. This can be improved by modifying surface residues with polyethylene glycol (PEG). Random derivatization usually creates an ill-defined mixture of molecules with varying degrees of PEGylation. A strategy to introduce PEG residues in a defined way is therefore much preferable over random modification. This is being tried using incorporation of unnatural amino acids with an alkyne side chain (Deiters et al. 2004). Subsequent modification with azide group-containing PEG residues generates proteins with site-specific PEGylation.
Many proteins are stabilized by disulfide cross-bridges. In recombinant proteins, these disulfide bonds are usually difficult to reproduce because of their redox sensitivity and the lack of relevant chaperones. A general method for the genetically programmed installation of directed redoxinsensitive cross-links has recently been devised (Neumann et al. 2010). The authors combined evolved orthogonal ribosomes with two pairs of mutually orthogonal tRNA/ aaRS pairs to incorporate two different unnatural amino acids into the same protein. The side chains of these amino acids contained an azide and an alkyne functional group that formed a cross-link upon addition of a Cu(I) catalyst.
Post-translational modifications (PTMs) are often present on proteins and can tremendously change their properties. However, recombinant proteins from E. coli are usually unmodified. It is therefore often necessary to use eukaryotic expression systems to produce correctly processed proteins, but even then the pattern of modifications is usually different from the natural situation. Lysine acetylation has been recognized as a widespread PTM in prokaryotes and eukaryotes (Choudhary et al. 2009;Kim et al. 2006;Zhang et al. 2009). The effect of the modification on protein function is largely unexplored. A method to genetically encode this PTM on recombinant proteins in E. coli has recently been devised (Neumann et al. 2008;Neumann et al. 2009). Hence, therapeutic proteins harbouring this modification can now be produced recombinantly without any knowledge of the modifying enzymes involved. Similar methods exist to produce proteins containing lysine methylation or tyrosine sulphation (Liu and Schultz 2006;Nguyen et al. 2009a). Other PTMs that cannot be encoded genetically to date, like glycosylations or ubiquitination, can be mimicked by incorporating a chemical handle into the protein at the site of the modification followed by in vitro derivatization (Liu et al. 2003).

Conclusion and perspective
In this review, we did not intend to give a complete list of all the parts available to a synthetic biology approach to pharmaceutical biotechnology and we apologize to those whose work was left unmentioned (please see Kiel et al. (2010), Ninfa (2010) and Topp and Gallivan (2010) for more recent reviews of this topic.) Our aim was to give a flavour of the potential impact that synthetic biology might have on the pharmaceutical industry. Future work should use these parts to create systems for the simultaneous production, identification and optimization of new drugs. Such an approach might look as follows: A reporter is engineered to detect a protein-protein interaction or an enzymatic activity of interest. To this end, one could develop biosensors that sense a reaction product or genetic circuits that are activated once a certain interaction is prevented. Such a sophisticated system will reduce the number of false hits during the screening process. This reporter system is subsequently combined with various in vivo compound libraries to isolate clones with the desired activity. Newly identified lead structures can be further improved afterwards, for example, with the help of unnatural amino acids. Their biosynthesis could be optimized for instance by using MAGE or by compartmentalizing the enzymes involved. Any improvement to the system could be identified or selected quickly by the reporter. A lot of work need to be invested before such an approach will become the mainstream of drug development. An important next step towards this aim would be to form well-characterized modular parts from individual components that can be freely and quickly combined in the true sense of a synthetic biology approach to pharmaceutical biotechnology. The automation of such an approach combined with the development of computational design tools for synthetic biology (Marchisio and Stelling 2009) will be an important technological advance that will help in its realization.


extfieldsofstudy: Chemistry Biology

text:
Dynamics of intracellular Ca$^{2+}$ oscillations in the presence of multisite Ca$^{2+}$-binding proteins

We study the dynamics of intracellular calcium oscillations in the presence of proteins that bind calcium on multiple sites and that are generally believed to act as passive calcium buffers in cells. We find that multisite calcium-binding proteins set a sharp threshold for calcium oscillations. Even with high concentrations of calcium-binding proteins, internal noise, which shows up spontaneously in cells in the process of calcium wave formation, can lead to self-oscillations. This produces oscillatory behaviors strikingly similar to those observed in real cells. In addition, for given intracellular concentrations of both calcium and calcium-binding proteins the regularity of these oscillations changes and reaches a maximum as a function noise variance, and the overall system dynamics displays stochastic coherence. We conclude that calcium-binding proteins may have an important and active role in cellular communication.

Animal cells communicate through the exchange of molecules secreted in the extracellular environment and/or by cell-cell contacts through the chemical interactions of molecules expressed at the cell surface. Cell communication is at the basis of animal physiology and pathology, and the information stored in the environment must ultimately be decoded in the cell and propagate to the nucleus where target genes can either switch on or off in response to these stimuli. Calcium (Ca 2+ ) is the most versatile second messenger in living cells and translates the information stored in the extracellular environment in time-dependent variations of Ca 2+ intracellular concentration. These may take the form of waves, bursts and oscillations that propagate in time and space in the cell and through adjacent cells [1,2,3].
Ca 2+ oscillations occur in a large number of cell types such as excitable (e.g. neurons, cardiac cells) and non excitable (e.g. hepatocytes, endothelial cells) cells, either spontaneously or after stimulation by hormones, cytokines and neurotransmitters, and they drive important functions such as brain and cardiac activity, immune cell activation, hormone secretion and cell death [1,2].
Two aspects of intracellular Ca 2+ oscillation have received little attention, with some notable exceptions [4,5]: Ca 2+ oscillations propagate in the cell in the presence of Ca 2+binding proteins that act as buffers, taking up to 99% of Ca 2+ in the cell; the mechanisms of Ca 2+ wave generation is intrinsically noisy. Here we address both aspects from a dynamical perspective.
Ca 2+ concentration within cells is controlled by reversible binding to specific classes of proteins that act as Ca 2+ sensors and decode the information carried by FM and AM modulation of the Ca 2+ oscillations [6]. Many intracellular proteins can bind Ca 2+ on multiple sites. For example, the activity of key enzymes such as CaM-kinase II is modulated either by direct binding to Ca 2+ or indirectly by chemical interaction with Ca 2+ -binding proteins [2]. Ca 2+ binding to proteins on multiple sites is not driven by enzymatic mechanisms and therefore its chemistry obeys to the law of mass action. In addition, the binding sites on Ca 2+ -binding proteins appear to be all equivalent in presence of the EF hand motif which allows the specific and reversible binding of Ca 2+ ions [2].
In this paper we investigate Ca 2+ dynamics in cells in the presence of Ca 2+ -binding proteins and noise. To this end, the vitamin D-dependent protein calbindin-D29K (CaL) has been chosen as an example of multisite Ca 2+ binding protein, representing a class of several Ca 2+ binding proteins that share many biochemical traits with CaL [2]. CaL is a protein with a molecular weight of 30 kDa, that is expressed in the cytosol of specific cells of the central nervous system such as hippocampal granule cells and cerebellar Purkinje cells [2]. CaL is a member of the EF hand superfamily Ca 2+ -binding proteins. To date, CaL has not yet been found to play a role in Ca 2+ -dependent regulation of enzyme activity although it is expected to act as an intracellular Ca 2+ buffer. This seemingly minor role may nonetheless have a profound effect on cell physiology, and indeed overexpression of CaL in cultured hippocampal pyramidal neurons affects synaptic plasticity and suppressed post-tetanic potentiation [10]. CaL has four reversible binding sites for Ca 2+ and, importantly, the on-off rates for the binding reaction have been determined experimentally. Experiments have also shown the presence of binding sites with two different affinities and a ratio of sites with high and low affinity of 3 to 1 or 2 to 2 [11].
We describe the dynamics of Ca 2+ reversible binding to CaL with the following differential system, which is formally the same that we used in a previous study of multisite protein modification ( [7,8]; see also [9] for a derivation of these equations): We still have to specify the function f ([Ca 2+ ], t); there are many different models of intracellular calcium oscillations, but this choice is not critical, and here we take the minimal and well-known model based on Ca 2+ -induced Ca 2+ release (CICR) as the basic model of Ca 2+ oscillations [12], which is sketched in fig.1 (For recent reviews, see also [3,6] and references therein).
The CICR model must be complemented by a stochastic term because the process of Ca 2+ wave generation is intrinsically noisy [13,14]. dynamics cannot be neglected, and we introduce a stochastic term (as in [15,17] that leads to the following stochastic differential system for the driving function f = f ([Ca 2+ ], t): where ] p ) and ξ(t) is Gaussian white noise with zero mean and variance D.
In these equations (see also fig.1), [Ca 2+ ] is the cytosolic Ca 2+ concentration, whereas [Y ] denotes Ca 2+ concentration in the IP3-insensitive intracellular store. v 0 = 10 −6 M s −1 is the input rate of Ca 2+ from the extracellular medium, v 1 = 7.3 · 10 −6 M s −1 is the parameter related to the IP3-modulated release of Ca 2+ from the IP3-sensitive store. V M 2 = whereas V M 3 = 500 · 10 −6 M s −1 is the maximum rate of release of Ca 2+ from that store into the cytosol in a process activated by cytosolic Ca 2+ ; K 2 = 10 −6 M, K R = 2 · 10 −6 M and K A = 0.9 · 10 −6 M are threshold constants for pumping, release and activation, respectively; k f = 1 s −1 is a rate constant that regulates the passive, linear leak of Ca 2+ from the IP3-insensitive store into the cytosol; k = 10 s −1 regulates the assumed linear transport of cytosolic Ca 2+ into the extracellular medium. The exponents n = m = 2 and p = 4 denote the Hill coefficients characterizing these processes.
The parameter β regulates the saturation of the IP3 receptor and acts as the control parameter which sets the level of the stimulus and varies from 0 to 1. We also introduce a parameter τ ≥ 1 s that scales the rates v 0 , V 1 , V M 2 , V M 3 , k f and k. In this way we can tune the Ca 2+ period of the CICR oscillator to match oscillations observed in real experiments.
We use equations (1) and (2) to investigate numerically the dynamic interplay between Ca 2+ , Ca 2+ -binding proteins and noise in the cell. We integrate the stochastic differential system with the Euler-Maruyama algorithm [16], and for stability reasons we take a short integration time step ∆t = 0.001s.
Before studying the influence of internal noise, we investigate the deterministic system's However, fluctuations of Ca 2+ concentration due to internal noise allow the system to cross the threshold again and oscillate even in the presence of supercritical CaL concentrations ( fig.3). One feature of the simulation outputs in fig.3 is that they are strikingly similar to time series sampled in real cells [18]. In these experiments, Ca 2+ spikes have been observed to occur randomly, and the average interspike interval T measured independently for hundreds of cells is correlated to the standard deviation σ in a rather broad range of T values [18]. We find the same result in our simulations ( fig.3).
Thus, both experiments and the numerical results shown in fig.3, suggest that noiseinduced Ca 2+ spiking and spiking periodicity depend on noise variance, and moreover the observed correlation between noise variance and oscillation frequency also suggest that what we are witnessing here is a form of stochastic coherence [17]. As in [17] we study this aspect using the regularity parameter R, which is the ratio between the average and standard deviation of the interspike interval (the "period" of the oscillations): As already noted in [17], the reciprocal of this quantity is just the "coefficient of variation" often used in neuroscience as an estimator of the regularity interspike intervals. In our simulations (see fig.4) we find that regularity peaks at a given noise variance, and that the position of the peak also depends on the CaL concentration. Thus the whole system dis-  [19]. We conclude that proteins, such as CaL, may not simply act as passive Ca 2+ buffers in cells, but be prime actors in the complex play of cellular communication. Rates v 0 , k and k f denote replenishment of the IP3-sensitive store with extracellular Ca 2+ , the efflux of cytosolic Ca 2+ from the cell and passive leak of Ca 2+ from the store Y into the cytosol. More details can be found in [3]. This classical scheme is modified here to take into account the action of Ca 2+ protein buffers such as calbindin (CaL) that binds Ca 2+ ions on multiple sites. Binding follows the law of mass action and is described by rates k on and k of f . Finally, in our model cytosolic Ca 2+ fluctuates because of internal noise.


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Recombinant Expression, Purification, and Functional Characterisation of Connective Tissue Growth Factor and Nephroblastoma-Overexpressed Protein

The CCN family of proteins, especially its prominent member, the Connective tissue growth factor (CTGF/CCN2) has been identified as a possible biomarker for the diagnosis of fibrotic diseases. As a downstream mediator of TGF-β1 signalling, it is involved in tissue scarring, stimulates interstitial deposition of extracellular matrix proteins, and promotes proliferation of several cell types. Another member of this family, the Nephroblastoma-Overexpressed protein (NOV/CCN3), has growth-inhibiting properties. First reports further suggest that these two CCN family members act opposite to each other in regulating extracellular matrix protein expression and reciprocally influence their own expression when over-expressed. We have established stable HEK and Flp-In-293 clones as productive sources for recombinant human CCN2/CTGF. In addition, we generated an adenoviral vector for recombinant expression of rat NOV and established protocols to purify large quantities of these CCN proteins. The identity of purified human CCN2/CTGF and rat CCN3/NOV was proven by In-gel digest followed by ESI-TOF/MS mass spectrometry. The biological activity of purified proteins was demonstrated using a Smad3-sensitive reporter gene and BrdU proliferation assay in permanent cell line EA•hy 926 cells. We further demonstrate for the first time that both recombinant CCN proteins are N-glycosylated.

Introduction
Fibrosis is characterised by the deposition of the extracellular matrix (ECM) proteins including collagens, structural glycoproteins, sulphated proteoglycans and hyaluronan into the subendothelial space of Disse [1]. The biomechanical and biochemical rebuilding of the tissue is induced by several important fibrogenic cytokines including transforming growth factor (TGF-b, platelet derived growth factor-B and -D (PDGF-BB, PDGF-DD), insulinlike growth factor-1 (IGF-1) and endothelin-1 (ET-1) [2]. TGF-b assists the wound repair and ECM deposition through intracellular Smad proteins that transfer signals from the cell membrane to the nucleus [3] and is of fundamental importance for the development of excess fibrous connective tissue [4]. In this process, the Connective Tissue Growth Factor (CCN2/CTGF) appears as the downstream modulator of TGF-bsignalling. In liver for example, CCN2/CTGF expression is in vitro triggered in hepatocytes, fibroblasts and other cell types after stimulation with TGF-b [5,6]. In vivo it could be shown, that the expression of CCN2/CTGF was persistent after TGF-b stimulation. It acts as a cofactor with TGF-b to induce hepatic fibrogenesis but is not considered to be a potent fibrogenic agent on its own [7]. However, one of its important functions is to stimulate the expression of ECM and proliferation of connective tissue cells to close wounds inside of internal organ and skin tissue [8][9][10].
CCN2/CTGF is one of the six members of the CCN protein family. The name of the family is an acronym of the first three identified members, namely CYR61 (CCN1), CTGF (CCN2) and the Nephroblastoma-overexpressed gene (NOV, CCN3). The other members are WISP1 (ELM1, CCN4), WISP2 (COP-1, CCN5) and WISP3 (CCN6). The different CCN proteins have a similar predicted secondary structure consisting of 5 individual modular domains that includes an insulin-like growth factor binding domain, a von Willebrand factor type C repeat, a thrombospondin type 1 repeat and a carboxyl-terminal cysteine knot [11,12]. Moreover, the CCN proteins show a high sequence identity between species with a highly conserved intramolecular network of cysteine residues [13]. They own a variety of biological functions that is strongly dependent on their cellular microenvironment [14] and fine-tuned by the activity and interplay of the individual structural domains [15].
CCN2/CTGF is a 38-kDa protein was originally identified as a growth factor secreted by human vascular endothelial cells [16] and defined as the chondrocyte-specific gene product [17,18]. It is over-expressed in many fibrotic disorders including scleroderma, kidney and liver fibrosis [19][20][21]12] and constitutive expression of CCN2/CTGF in tissues is considered to indicative for pathological lesions [5]. CCN2/CTGF promotes endothelial proliferation, migration, survival and adhesion in angiogenesis [22]. Furthermore, CCN2/CTGF acts as an immediate early gene that are induced by serum or growth factors and believed to play important roles in the control of cell proliferation [9]. The TGF-b dependency of CCN2/CTGF is mainly controlled by sequence elements within the promoter that include a Smad-binding site and a defined TGF-b response element [20,23].
Another member of the CCN family is the CCN3/NOV protein that under certain conditions counteracts CCN2/CTGF functions [24,25]. CCN3/NOV was first identified in Myeloblastosis-associated virus-1-induced avian nephroblastomas [26,27]. It is over-expressed in a broad range of tumors including Wilm's tumor and is further associated with neovascularisation and vascular injuries in which it promotes pro-angiogenic activities through integrin receptors in vascular endothelial cells [28,29]. Some other studies have shown that CCN3/NOV inhibits glioma cell and PDGF-dependent glomerular cell proliferation [30,31].
However, the molecular understanding of CCN2/CTGF and CCN3/NOV is still at the beginning and several key questions are still unanswered. In particular, data is missing that explains the regulatory network in which CCN2/CTGF promotes TGF-b action that is necessary to maximally induce certain genes including collagen type I and a-smooth muscle actin. Moreover, the mechanisms that are required for the reported Yin/Yang regulation of CCN2/CTGF and CCN3/NOV are still an enigma. Presently, the performance of studies addressing these mechanistic issues are somewhat limited by the lack of larger quantities of biological active of respective CCN proteins.
In the present study, we aimed to develop devices that allow the production of biological active recombinant CCN2/CTGF and CCN3/NOV. We report about the establishment of transgenic cell lines that over-express human CCN2/CTGF and an adenoviral expression vector for production of rat CCN3/NOV. We have established protocols for purification of these two CCN, and biochemically and biologically characterised the recombinant proteins by mass spectrometry, tryptic in-gel-digests, and proliferation and gene reporter assays in EANhy 926 cells. Moreover, we demonstrate for the first time that both CCN2/CTGF and CCN3/ NOV are N-glycosylated and that the predicted Yin/Yang activity both CCN proteins can be demonstrated in EANhy 926 cells.

Transfection experiments
All transfections were done with the FuGENE 6 transfection reagent (Roche) according to the manufacturer's instructions. Briefly, HEK cells were seeded at 2610 5 cells per 6-well plate (Sarstedt, Nürnbrecht, Germany) and incubated overnight in culture medium. The medium was renewed and cells were cultured for another 2 h period prior transfection. Sixteen hours later, the medium was changed and clones were selected with indicated antibiotics.

Stable expression of human CCN2/CTGF in HEK cells and in the Flp-In-293 systems
The full length human CCN2/CTGF cDNA sequence was amplified from expression clone pCEP4-hCTGF [34] that was kindly provided by W. Sebald (Department of Physiological Chemistry II, Theodor-Boveri Institute for Life Sciences (Biocenter), University of Wuerzburg, Germany) using primers hCTGFfor (59-GAA TTC ATG ACC GCC GCC AGT ATG GG-39) and hCTGFrev(59-CTC GAG TGC CAT GTC TCC GTA CAT CT-39). The amplicon was cloned into the pGEM TM -Easy cloning vector (Promega, Mannheim, Germany). Subsequently, the cDNA was released by the NotI endonuclease and subcloned into the multiple cloning site of vector pcDNA5/FRT/TO (Invitrogen) designed for use with the Flp-In-293 system that allows production of stable cell clones expressing the gene/protein of interest. The correct orientation within vector pcDNA5/FRT/TO-hCTGF was verified by sequencing. This vector system contains a Hygromycin B resistance gene cassette allowing selection and maintenance of transgenic clones. For generation of stable expression clones, cells (HEK293A or Flp-In-293) were seeded on 6-well plates and transfected with pcDNA5/FRT/TO-hCTGF alone (HEK293A) or pcDNA5/FRT/TO-hCTGF and pOG44 (Invitrogen) (Flp-In-293) containing the transferase/integrase gene. Transfected cells were selected with increasing concentrations of Hygromycin B ranging from 50-400 mg/ml and dying of untransfected cells was visualized by light microscopy. The ideal concentration was determined to 150 to 200 mg/ml of Hygromycin B for both cell lines. Individual cell clones were established by limiting dilution (0.5 cells/well in a 96-well plate) and numbered consecutively (HEK clones: 1/1, 1/2, 1/3 etc.; Flp-In-293 clones: WB1, WB2, WB3 etc.). Expression and secretion of recombinant CCN2/CTGF was analyzed in ELISA [35] and Western blot using the CCN2/ CTGF-specific antibody sc-14939 (Santa Cruz). Stable cell clones were frozen at 280uC using standard cryoconservation protocols. Five of 120 initial cell clones were further expanded for production of recombinant CCN2/CTGF. The expression of recombinant CCN2/CTGF in each of both systems (HEK, Flp-In-293) was documented for over 100 passages.

Immunocytochemistry
HEK and Flp-In-293 cells were seeded on poly-L-lysine-coated glass cover slips that were prepared following a standard protocol.
On the next day, the medium was renewed and cells were incubated for further 24 hr. Then the cells were washed in PBS, fixed in 4% (v/v) paraformaldehyde in 16 PBS (pH 7.4) for 10 min, and permealized for 2 min on ice in 0.1% (w/v) sodium citrate and 0.1% (v/v) Triton X. Cells were blocked for 10 min at RT in Biotin Blocking Reagent (X0590) obtained from DAKO (Hamburg, Germany) following manufacturer's instructions. Unspecific binding sites were further blocked in 50% (v/v) FCS, 1% BSA (w/v), and 0.1% (v/v) fish gelatine in PBS (pH 7.4) for 1 h. Finally, the cells were incubated overnight at 4uC in a 1:300 dilution of the anti-human CCN2/CTGF antibody (sc-14939) in PBS containing 1% (w/v) BSA. Cells were washed and incubated with an anti-goat-biotinylated antibody (1:300, DAKO) for 1 h, and further with a Streptavidin/FITC-conjugate for 30 min at RT. The nuclei were stained with DAPI. As a control, the cells were incubated with an unspecific immunoglobulin.

Western blot analysis
The cells were washed three times in PBS and lysed in 250 ml RIPA buffer (50 mM Tris/HCl (pH 7.4), 1% (v/v) NP-40, 0.25% (w/v) sodium deochycholate, 150 mM NaCl, 0.1% (w/v) SDS, 1 mM EDTA, 1 mM PMSF, 1 mM Na 3 VO 4 , and 1 mM NaF) per well of the 6-well plate. The protein concentration was measured using the Micro BCA protein assay (Pierce Thermo Scientific, Bonn, Germany). Proteins were separated in Bis/Tris 4-12% SDS-PAGE (Invitrogen) in MES buffer and transferred to nitrocellulose membranes (Schleicher & Schüll, Dassel, Germany). For documentation of equal protein loading, the membranes were stained with Ponceau S and blocked for 30 min in 5% non-fat dried milk dissolved in TBST containing 10 mM Tris (pH 8.0), 150 mM NaCl, and 0.1% (v/v) Tween-20. The membranes were incubated overnight with a goat-anti-CCN2/CTGF (sc-14939) at 4uC under permanent shaking. The membranes were rinsed three times in TBST and incubated with a monoclonal donkey antigoat-HRP conjugate (sc-2056, Santa Cruz) at RT for 1 h and signal detection was conducted by use of the SuperSignal West Pico Chemiluminescent Substrate (Pierce).

Generation of adenoviral expression vector AdEasy-CMV-rNOV
To subclone the coding region of rat CCN3/NOV, a PCR was performed using plasmid clone IRAKp961P24175Q (ImaGENE, Berlin, Germany) as a template and primers 59-(TTG TAG AAT TCA GCA GGC AGA ACA TG)-39 and 59-(TTA CCG GTA CAT TTC TCC TCT GCT)-3. The amplicon was first cloned into pGEM-T-Easy. Subsequently, the NOV containing sequence was released by EcoRI/AgeI and cloned into respective sites of vector pcDNA3.1/V5-HisA. Thereafter, the CCN3/NOV expression cassette was cut out by digest with EcoRI and PmeI, blunted by Klenow, and cloned into vector adenoviral shuttle vector pShuttle-CMV (Stratagene, Agilent Technologies, Waldbronn, Germany). For generation of a recombinant adenoviral plasmid, the resulting vector pShuttle-CMV-rNOV was linearized by PmeI digest and cotransfected with adenoviral backbone vector pAdEasy-1 (Stratagene) into electroporation competent BJ5183 bacteria. A recombinant vector pAdEasy-1-CMV-rNOV was isolated, characterized by restriction digest and sequencing, and transfected into 293A cells to prepare adenoviral particles using standard transfection, amplification and purification techniques.

Purification of recombinant CCN2/CTGF
For production of recombinant human CTGF, transgenic cells were seeded at 2610 6 cells/culture dish and cultivated in growth medium for 24 h. The medium was replaced one day later with medium without FCS and expression/secretion of CCN2/CTGF was allowed for 48 h. For purification of CCN2/CTGF, we followed a two-step protocol. In the first step (heparin affinity chromatography), we took advantage of the ability of CCN2/ CTGF to bind to heparin. The enriched growth medium was centrifuged for 20 min at 50006g and supernatant was filtered (0.22 mm). Then the growth medium was equilibrated with 1/10 VT of 100 mM Tris/HCl (pH 7.0) and subjected with a syringe to a HiTrap TM HP Heparin column (5 ml) that was equilibrated with 25 ml of 10 mM Tris/HCl (pH 7.0). The column was then washed with 25 ml 10 mM Tris/HCl (pH 7.0) and the recombinant protein was stepwise eluted from the heparin matrix with 10 mM Tris/HCl (pH 7.0) containing increasing concentrations of NaCl ( . The purest CCN2/CTGF fraction was eluted at 0.7 M NaCl. The content of each fraction was tested by Western blot or ELISA. CTGF containing fractions were pooled and subjected to a second cycle of heparin affinity chromatography. Then the 0.7 M NaCl fraction of the second Heparin affinity chromatography was applied to a Superdex Sepharose 16/60 size-exclusion chromatography that was run on a FPLC chromatographic system from GE Biosciences equipped with a P-900 photometer. Fractions of 3 ml were collected and tested for the content of recombinant CCN2/CTGF by ELISA or Western Blot. The purity of protein solution was evaluated by SDS-PAGE and Coomassie Brilliant Blue staining. All buffers and solutions used during the purification of rhCTGF was filtered (0.22 mm) filtrated and kept at 4uC.

Purification of recombinant CCN3/NOV
The purification of recombinant rat NOV was performed from pooled supernatants of COS-7 cells that were infected with the adenoviral vector Ad-CMV-NOV and later harbored in DMEM without FCS 48 h. After centrifugation (20 min, 50006g) and filtration (0.22 mm), the equilibrated supernatant (10 mM Tris, pH 7.0) was subjected to heparin affinity chromatography. After washing (25 ml of 10 mM Tris/HCl pH 7.0 and 25 ml of 10 mM Tris/HCl pH 7.0/0.15 M NaCl), recombinant NOV was eluted with 0.8 M NaCl in 10 mM Tris/HCl (pH 7.0). In a second step, NOV containing fractions were pooled and further purified on a HiLoad 16/60 Superdex TM column with 10 mM Tris/HCl (pH 7.0) as running buffer.

CCN2/CTGF ELISA
The ELISA for quantification of human CCN2/CTGF was described before [35]. Briefly, the antibodies (rabbit-anti-human CCN2/CTGF (sc-25440) for plate coating, goat-anti human CCN2/CTGF (sc-14939) for detection, and donkey anti-goat IgG-HRP (sc-2056) for visualisation) were all obtained from Santa Cruz. The microtiterplates were first coated overnight at 4uC with a 1:1000 dilution of antibody sc-25440 in coating buffer [50 mM natrium carbonate (pH 9.6): 1.59 g/l Na 2 CO 3 and 2.93 g/l NaHCO 3 ]. The plates were then rinsed three times with PBST (PBS with 0.05% Tween-20) and supernatants were mixed with an equal volume of PBST and dispensed into the wells and incubated at RT under shaking for 2 h. The probes removed, wells rinsed with PBST and the second antibody against CCN2/CTGF (sc-14939) dispersed in a 1:1000 dilution into every well and incubated for 2 h at RT. After a further washing, the anti-goat IgG-HRP conjugate was dispersed in a 1:15000 dilution for 1 h. The plates were washed three times and developed by addition of the 3,39,5,5-tetramethyl benzidine (TMB) substrate at RT for 10 min. The reaction was stopped in 2 M H 2 SO 4 and the absorbance at 450 nm was determined in a Wallac Victor 1420 Multilabel counter. The concentration was measured in duplicate.

2-D PAGE and mass-spectrometric analysis
Immobilized pH gradient strips (ReadyStrip TM IPG Strip, pH range 3-10, #163-2000, Bio-Rad Laboratories, Philadelphia, PA) were re-hydrated in the PROTEANH IEF focusing tray. The protein sample was re-suspended in 125 ml of re-hydration buffer (9 M Urea, 4% CHAPS, 0.4% Ampholyte, 65 mM DTT and Bromphenol Blue) and applied as a line along the edge of channel in the PROTEANH IEF focusing tray and covered with the immobilized pH gradient (IPG) strips (7 cm length). Before starting the active re-hydration, the IPG strip was passively rehydrated for one hour until the liquid was absorbed by the gel bed. The IPG strips with the re-hydration buffer/protein solution were covered with mineral oil to prevent the evaporation during active re-hydration and entrance of the proteins into the gel of the IPG strip overnight. Wet electrode wicks were inserted between the IPG strips and the electrodes. The active re-hydration of the protein species on the IPG strips was done in the PROTEANH IEF Cell overnight (16 hr): 50 V, 20uC. The focussing program was performed in the PROTEANH IEF Cell (Bio-Rad). The first step of focusing was done rapidly linear at 500 V for 250 Vh, the second step was done from rapidly linear at 1000 V for 500 Vh. The third step was done rapidly linear at 8000 V for 6500 Vh. The program ended at constant 500 V.
Before running the second dimension, the IPG strips were equilibrated in SDS-containing buffers. The first equilibration buffer containing DTT reduces sulfhydryl groups and the second equilibration buffer containing iodoacetamide alkylates the reduced sulfhydryl groups. Briefly, the strips were shortly washed with Milli-Q water and soaked for 15 min in a buffer containing 6 M Urea, 30% (v/v) glycerol, 2% (w/v) SDS, 0.065 M DTT, and 0.05 M Tris (pH 8.8) on an orbital shaker. Then, the strips were shortly washed in Milli-Q water and equilibrated for 15 min in a buffer containing 6 M urea, 30% (v/v) glycerol, 2% (w/v) SDS, 0.135 M iodoacetamide, and 0.05 M Tris (pH 8.8) on the orbital shaker. The equilibrated IPG strips were placed on the top of the 12% SDS-PAGE that was run in Tris/glycine buffer (248 mM Tris, 192 mM glycine, 1% (w/v) SDS, pH 8.3) for 20 min at 120 V to force the immobilized protein spots from the strip. Subsequently, the separation of the protein spots was performed for 40 min at 180 V. The final gels were analysed by colloid Coomassie staining, Western blot and in gel digestion was realized prior to ESI-MS/MS analysis.

ESI/TOF-MS/MS
The identity of CCN3/NOV and CCN2/CTGF was verified by MS/MS-identification of individual fragments obtained by tryptic digestion. The polyacrylamide gel was rinsed two times in Milli-Q water to remove excess SDS from the gel, reducing strong background during sensitive Coomassie blue staining [36]. Then the gels were stained overnight in 5% (w/v) aluminium sulphate hexadecahydrate (06421; FLUKA, Sigma-Aldrich, Munich, Germany), 10% (v/v) ethanol, 0.02% (w/v) Coomassie Brilliant Blue (9598.1, CBB-G250, Roth) and 2% (v/v) of 85% ophosphoric acid (79620, FLUKA). Thereafter, the gels were rinsed 2 times in Milli-Q water and bands of interest were excised with a clean scalpel avoiding background gel volume around of the

MALDI-TOF mass spectrometry
Desalting of peptides was done by using the ZipTipC m-C8 devices according to the manufacturer's instruction (Millipore Corporation, Billerica, MA). The dried protein samples were resuspended in 10 ml of 5% (v/v) formic acid in HPLC pure water. The ZipTipC m-C8 was pre-wetted in 25% (v/v) of mixture of 5% (v/v) formic acid in HPLC pure water and 75% (v/v) acetonitrile and equilibrated with 5 volumes of 5% (v/v) formic acid in HPLC pure water. The proteins were applied on the ZipTipC m-C8 in 10 aspiration and dispensing cycles. Then, the surface of the column of the ZipTipC m-C18 was washed with 5 volumes of 5% (v/v) formic acid in HPLC pure water and the proteins were eluted into a clean vial by dispensing of 2-4 ml of elution solution containing 25% (v/v) of a 5% (v/v) aqueous formic acid solution in HPLC pure water and 75% (v/v) acetonitrile. The solution was aspirated and dispensed through the ZipTipC m-C8 at least 10 times. Subsequently, 1 ml of the eluted protein was dried on the 96well MALDI target plate (Gilson, Middleton, USA) and 1 ml of the Matrix solution containing sinapinic acid in 30-50% of acetonitrile, 0.1% trifluoracetic acid and HPLC pure water was pipetted on the dried protein drop. The mass of recombinant CTGF and NOV were determined with the MALDI-TOF/TOF mass spectrometer method (ultrafleXtreme MALDI-TOF/TOF mass spectrometer, Bruker Daltonics, Bremen, Germany). The protein standard II (Bruker) that contains three standard proteins (Trypsinogen, Protein A, Bovine Serum Albumin) with average m/z 23982, 44613, and ,66500 was used for testing and calibration of the MALDI mass spectrometer.
Biological activity of purified CCN proteins (i) Cell proliferation. The biological activity of purified recombinant CCN2/CTGF was tested om EANhy 296 cells. Therefore, 3610 4 cells were seeded in the wells of a 12-well plate and cultured overnight in growth medium. The next day, the medium was changed to starvation medium (0% FCS, 1 mg/ml BSA) and cells were serum-starved for 16 hr. Thereafter, the medium was renewed, different amounts of recombinant CCN2/ CTGF (100 ng to 1.5 mg/ml) added, and cells incubated for a further 24 hr period. Then the medium was replaced against labeling medium (DMEM, 1 mg/ml BSA, and 0.01 mM 5bromo-29-deoxyuridine) and cells incubated for a further 24 hr  period. Finally, the incorporation of BrdU was quantified with the 1-Step TM BrdU ELISA (Roche) according to the manufacturer's instructions. The absorption was measured in a Wallac Victor 1420 Multilabel counter. (ii) Luciferase assay. EANhy 926 cells were seeded with 1610 5 cells/well in 6-well plates and incubated overnight. The next day the medium was renewed and transfected with 1 mg of the (CAGA) 12 -MLP-Luc vector using 3 ml of the TransIT-LT1 transfection reagent (Mirus, VWR International, Darmstadt, Germany) after another 2-hr incubation time. The next day, the serum content of the medium was reduced to 0.5% FCS and cells starved for 16 hr. Thereafter, cells were stimulated in medium (0.2% FCS) with indicated concentrations of cytokines for 24 hr. Finally, cells were lysed in 16 luciferase lysis buffer (Promega), LAR substrate added, and the luciferase activity measured in a Wallac Multilabel counter. The luciferase activity given in light counts per second was normalized against the protein concentration in each sample and given as relative light units (RLU).

Purification of recombinant human CCN2/CTGF and recombinant rat CCN3/NOV
We cloned expression constructs that direct the expression of full-length human CCN2/CTGF and rat CCN3/NOV with short fusions at their carboxyl-terminal regions ( Figure S1). For overexpression of human CCN2/CTGF, we established stable transfected HEK293A (HEK 1/1, HEK2/2) and Flp-In-293 cell clones (WB4) that show a strong expression of respective protein ( Figure 1A-1C) that is observable even after 100 cell passages ( Figure S2). In addition, we established an adenoviral expression vector that directs high expression of rat CCN3/NOV ( Figure 1D). Based on their property to bind heparin, we purified respective CCN proteins by use of affinity chromatography (Figure 2). Protein quantification was done with a standard protein quantification assay and specific measurement using in-house-CCN2/ CTGF [35] and in-house CCN3/NOV ELISAs (Borkham-Kamphorst and Weiskirchen, unpublished) as well as Western blotting. In addition, the purity of respective recombinant CCN proteins was analysed by applying 1 mg of recombinant proteins to 4-12% Bis/Tris SDS-PAGE electrophoresis following Coomassie stain. In this analysis, recombinant human CCN2/CTGF typically appeared as two distinct bands in the range of 38-40 kDa, while rat CCN3/NOV appeared as one broad smeary band with a molecular weight of ,48 kDa. When necessary, we further purified the proteins by Sepharose size-exclusion chromatography ( Figure 3). Interestingly, the buffer salt content showed a strong influence on the elution profile of recombinant CCN proteins. Affinity purified CCN3/NOV was fractioned with a bipartite chromatogram when the gel filtration was run in a buffer that simply contained 10 mM Tris ( Figure 3A). While commercially available CCN2/CTGF that was expressed in bacteria showed a tripartite elution profile ( Figure 3B), the protein that we have purified from cell supernatants showed a distinct peak at very low molecular weight ( Figure 3C). However, when the same protein was run in the same buffer with addition of 150 mM NaCl, we found that the peak was shifted to a range at which high molecular weight proteins were eluted ( Figure 3D).
We next tested the stability of purified recombinant CCN3/ NOV and CCN2/CTGF. Therefore, we analysed respective protein solutions that were stored at 4uC and 280uC for up to three months by SDS-PAGE and Coomassie stain under reducing and non-reducing conditions (Figure 4). This analysis revealed that both proteins were rather stable and showed no significant degradation after storage. When determined the total mass of purified human CCN2/CTGF and rat CCN3/NOV by MALDI-TOF, we observed average m/z values (CCN2/CTGF, 38269 Da; CCN3/ NOV, 44993 Da) that were roughly in the range of our expectation ( Figure 5). However, compared to our calibration proteins (Trypsinogen, Protein A, Albumin) that produced narrow peaks ( Figure 5A), the recombinant proteins produced broad peaks ( Figure 5B, and 5C) in this analysis that were not really appropriate to determine the mass. To characterise this phenomenon in more detail, we next performed 2D-gelelectrophoresis in which the proteins were separated on IPG strips according to their isoelectric points (1D) and subsequently by their size using 10% SDS-PAGE (2D). Again, the recombinant proteins produced smears in these 2D gels ( Figure 6) that were completely immunoreactive to specific antibodies suggesting that the complete smears resulted from different biochemical subspecies. We isolated spots/bands from 1D-(CCN2/CTGF) or 2D-gels (CCN3/NOV), subjected them to trypsin in-gel digesting and identified resulting peptides by ESI-TOF/MS and subsequent analysis using the MASCOT search engine (e.g. Table S1 and Table S2) for rapid protein identification using mass spectrometry data [37]. In this analysis, the sequence coverage was 34% for human CCN2/CTGF and 26% for rat CCN3/NOV. Subsequently, we demonstrated the biological activity of the purified proteins. The biological activity of recombinant CCN2/ CTGF was proven by stimulation of endothelial cell line EANhy 926 that represents a human intraspecies hybrid [33] that was previously tested to be highly sensitive against CCN2/CTGF [34]. When cells were stimulated for 24 h with different concentrations of purified CCN2/CTGF (100-2000 ng/ml) in serum-free medium, we observed a dosis-dependent increase of proliferation, while bacterial expressed CCN2/CTGF (bCTGF) that is commercially showed no stimulatory activity in this proliferation assay (Figure 7). Recombinant CCN2/CTGF induced a 1.6 fold BrdU incorporation compared to unstimulated controls. PDGF-BB (25 ng/ml) and TGF-b1 (1 ng/ml) that were incorporated in this assay as positive and negative controls induced a 2.9-fold induction or a 50% suppression of BrdU incorporation. Purified rat CCN3/ NOV showed no effect on the BrdU incorporation in this assay (data not shown).
To analyse if CCN2/CTGF interferes with TGF-b signalling, we performed reporter gene assays in with EANhy 926 cells using a reporter containing Smad3/Smad4 binding sequences, termed CAGA boxes, that were originally derived from the promoter of the human PAI-1 gene and is a read-out-system for phosphorylated Smad3 [32]. We transfected EANhy 926 cells with the reporter (CAGA) 12 -MLP-Luc vector and stimulated the cells in serum-free DMEM for 24 hr with different concentrations/ combinations of recombinant CCN2/CTGF, CCN3/NOV, TGF-b1, PDGF, bCTGF, and an antagonizing antibody (L-20) directed against CCN2/CTGF (Figure 8). The obtained luciferase activities were normalised to the protein amount and calculated as light count per second per mg protein (lcps/mg protein) and relative luciferase units (RLU) were compared to un-stimulated controls that were set to 1. In this assay, TGF-b1 (1 ng/ml) stimulation leads to a 6.2-fold increase, while PDGF-BB (50 ng/ml) stimulation decreases luciferase activity to 0.18 of the unstimulated control. Stimulation with 2 mg/ml of purified CCN2/CTGF induced the luciferase activity by a factor of 10, while the stimulation with 1 mg/ml of purified NOV decreased the luciferase activity to 0.6 fold. In the presence of the capturing antibody, the stimulatory effect of CCN2/CTGF on luciferase activity was negotiated. In line, the capturing antibody itself decreases luciferase activity to 0.4-fold of unstimulated control suggesting that the biological activity of endogenous expression of CCN2/ CTGF in EANhy 926 cells ( Figure S3) is blocked by sequestering. There was no increase of luciferase activity observable in the bCTGF-stimulated and transiently transfected EANhy 926 cells. We could show a significant difference between stimulated (500 ng/ml, 1000 ng/ml) and unstimulated EANhy 926 cells (p#0.05) using double-sided t-test ( Figure S4).
We next tested the effect of the purified proteins on biochemical cell activity. Therefore, equal cell numbers of EANhy 926 cells were seeded, serum-starved overnight and stimulated with TGF-b1, PDGF-BB, recombinant human CCN2/CTGF and rat CCN3/  NOV for 24 hr. After the incubation period, the cells were washed and whole cell lysates for comparative quantification (the amount of proteins in unstimulated cells were set to 1) of total protein concentration prepared. We found that cells treated with PDGF-BB had a 1.6-fold higher protein amount than unstimulated controls. In EANhy 926 cells that were stimulated with CCN3/ NOV (500 ng/ml), the protein amount was decreased to 0.6-fold of the unstimulated control. Likewise, stimulation with 1 ng/ml of  TGF-b1 decreased the protein amount to 0.75-fold of the unstimulated control. CCN2/CTGF had no influence on the protein amount (Figure 9).
Based on the finding that the purified CCN proteins appeared not as unique populations in gel electrophoresis (cf. Figure 6) and showed atypical chromatography behaviour (cf. Figure 3), we analysed if the heterogeneity resulted from glycosylation. When we analysed the human CCN2/CTGF and rat CCN3/NOV protein sequences for potential N-glycosylation sites using the NetNGlyc algorithm (http:// www.cbs.dtu.dk/services/NetNGlyc/), we found that both proteins carry a potential N-glycosylation site ( Figure S5). To test if the proteins are indeed N-glycosylated, we first analysed if the respective proteins have lectin binding capacity using a Con A-HRP conjugate ( Figure 10). As negative and positive controls, 1 mg BSA and the soluble highly glycosylated extracellular part of the PDGF-BB receptor [38] was incorporated into this assay. Additionally, bacterial expressed recombinant CTGF (bCTGF) that should contain no sugar groups was analysed in parallel. In this analysis, we found that both recombinant proteins from eukaryotic sources and the extracellular domain from the PDGF type b receptor (PDGFRb) were detectable with the Concanavalin (ConA)-HRP conjugate suggesting that these proteins carry sugar groups. In contrast, CCN2/ CTGF that originated from a bacterial source was only detected by the L-20 antibody but not by Con A. In a second set of experiments, we tried if the purified CCN proteins could be enzymatically deglycosylated by Endo H (removes only high mannose and some hybrid types of N-linked carbohydrates) or PNGase F (removes all types of N-linked carbohydrates). This analysis revealed that incubation of CTGF with one of both enzymes was suitable to remove the affinity for the Con A-HRP conjugate, while the respective affinity of CCN3/NOV was only removed after treatment with PNGase F (Figure 11). In addition, the treatment with PNGase F resulted in a marked alteration of electrophoretic mobility.
A previous study reports that CCN2/CTGF and CCN3/NOV can regulate their expression in an Yin/Yang manner and that either the addition or over-expression of CCN3/NOV produces a marked down-regulation of CCN2/CTGF in mesangial cells [25]. To test if this effect is also reproducible in EANhy 926 cells, equal cell numbers were seeded, serum-starved overnight, and cultured in conditioned medium enriched in recombinant human CCN2/ CTGF. Thereafter, the cells were washed, whole cell lysates prepared and analysed for CCN3/NOV expression revealing that EANhy 926 cells, that normally express both CCN proteins, produce less amounts of endogenous CCN3/NOV after CCN2/ CTGF treatment ( Figure 12).

Discussion
Protein studies aiming to unravel biological functions require pure and native proteins that might be further dependent on their Figure 11. Measurement of mitogenic activity of different growth factors. EANhy 926 cells were incubated with different concentrations of growth factors in serum-free medium. Protein amounts of whole cell lysates were measured by Micro BCA assay after 48 hr. In this analysis, the protein amount in untreated cells was set to 1. The experiments were repeated three times. Shown are the means 6 SD of one representative experiment that was performed in triplicate. doi:10.1371/journal.pone.0016000.g011 correct post translational modifications. It is known that the correct configuration of carbohydrate chains is important in the signalling of growth factors [39].
In this study, we have established novel devices for expression of recombinant human CCN2/CTGF and rat CCN3/NOV (Figure 1). By use of heparin affinity and size exclusion chromatography, we were able to purify both CCN proteins to heterogeneity ( Figure 2). Typically, these expression devices allow to purify 200-300 mg of recombinant human CCN2/CTGF from 400 ml growth medium of stable transfected Flp-In-293 clones and approximately 300 mg recombinant rat CCN3/NOV from 400 ml conditioned supernatant of COS-7 cells that were infected with Ad5-CMV-rNOV. The identity of purified proteins was demonstrated by mass spectrometry ( Figure 5) and trypsin in-gel-digest ( Figure 6) following identification of resulting peptides by ESI-MS/MS and subsequent analysis using the MASCOT algorithm.
We further demonstrate that the purified proteins are highly stable when stored at 280uC or 4uC (Figure 4). Interestingly, the retention time of purified CTGF in size exclusion chromatography was strongly dependent on the salt content of chosen buffer. Low salt concentration prolonged the retention time, while higher salt concentrations (150 mM) leads to shorter retention times ( Figure 3). Although this phenomenon is known for many other proteins, the drastic shift of the elution time might indicate that CCN2/CTGF strongly interacts with the sepharose matrix representing a crosslinked, beaded-form of a polysaccharide polymer material (agarose) that is crosslinked through lysine side chains.
The biological activity of the isolated proteins is the key indicator of successful expression. For recombinant CCN2/CTGF we have shown the biological activity by use of its positive influence on proliferation in EANhy 926 cells [34] and in a reporter gene assays using the Smad3/4-sensitve reporter (i.e. (CAGA) 12 -MLP-Luc)) that originally originated from the human PAI-1 gene promoter [32]. After addition of CCN2/CTGF, we found an increase of proliferation in BrdU incorporation assay of around 150% (Figure 7), values that were similar to those observed in pancreatic stellate cells [40]. In this assay, the EANhy 926 cells were also increased after stimulation with PDGF-BB and decreased after TGF-b1 stimulation and moreover bacterial expressed CCN2/CTGF failed to influence proliferation of EANhy 926 cells. The results obtained after transient transfection of EANhy 926 cells with the reporter vector (CAGA) 12 -MLP-Luc showed an increase of the firefly luciferase read that was dose-dependent and blocked by the addition of an antagonistic antibody (Figure 8). Even more, this antibody was suitable to block the stimulatory effect of endogenous CCN2/CTGF. Contrarily, the stimulation with recombinant NOV leads to a 0.6-fold decrease in luciferase activity suggesting that NOV might evolve opposing activities towards CCN2/CTGF. That finding fits well with the recent hypothesis that the expression of CCN2/CTGF and CCN3/NOV is controlled in an Yin/Yang manner [25] and was further highlighted by the fact that the expression of CCN3/NOV in EANhy 926 cells could be decreased by supplementation of growth medium with CTGF ( Figure 12). The most interesting finding of this study is the demonstration that both purified recombinant CCN proteins are N-glycosylated (Figures 10 and 11). Both proteins (i) have affinity for the Con A-HRP conjugate that is lost after treatment with endoglycosidases, (ii) migrate as diffuse bands in SDS-PAGE, and (iii) appear not uniform in mass spectrometry. We could demonstrate a prominent shift in molecular weight after deglycosylation with PNGase F for CCN3/NOV in SDS gel electrophoresis that is associated with a reduced lectin binding activity. Likewise CCN2/ CTGF that originated from eukaryotic sources had affinity for the Con A-conjugate that was not found in CCN2/CTGF that was expressed in bacteria and lost after treatment with either PNGase F or Endo H (Figure 11). Prediction that have been done on the ExPASy Proteomics Server show two N-glycosylation sites at positions 28 NCSG (high potential) and 225 NASC (low potential) in human CCN2/CTGF and at positions 91 NETG (potential) and 274 NCTS (low potential) in rat NOV. There were no predictions for O-glycosylation sites in human CCN2/CTGF and only one predicted O-glycosylation site at threonine 43. The activity of growth factors and signalling proteins are depended on the post-translational modifications including glycosylation and correct building of the intramolecular disulfide bond network. These modifications are difficult to perform in E. coli expressing system [41] but obviously accurately done in our eukaryotic expression systems (i.e. HEK, COS-7). Although we do not yet know if N-glycosylation is a post-translational modification that is important for the folding or biological activity of these CCN proteins, it might be an essential prerequisite for proper protein secretion and adhesion/incorporation into the ECM or to prevent premature degradation.
Incubation of EANhy 926 cells with conditioned medium isolated from WB4 over-expressing human CCN2/CTGF resulted in an increase of endogenous CTGF and suppression of endogenous CCN3/NOV. This data confirms previous suggestions that exogenous CCN2/CTGF is able to suppress expression of CCN3/NOV in a Yin/Yang manner [25].
In summary, we have established an expression and purification system to manufacture biological active recombinant human CCN2/CTGF and rat CCN3/NOV in permanent mammalian cell lines. Both CCN proteins are rather stable and can be stored at 4uC or 280uC for at least 3 month without disintegration. The biological activity of both proteins was demonstrated in gene reporter and proliferation assays.
The opportunity to prepare large quantities of biological active CCN2/CTGF and CCN3/NOV proteins will now allow us to address key questions of these CCN proteins in regard to structure, function, post-translational modifications (e. g. glycosylation), involvement in individual signal cascades, and further provide a foundation to address their therapeutic potential in various disease models.