Introduction
Understanding the intricate organisation and interaction of organisms, organelles, tissues, cells, and biomolecules, and how their dysregulation results in disease, is vital for the effective development of potential therapeutics and experimental platforms. The central dogma of molecular biology provides the fundamental building blocks of almost all eukaryotic functional pathways, through the relationship of DNA (transcription), RNA (translation) and protein (expression).
It is therefore no wonder that the application of molecular biology is essential in the development of new laboratory models, in understanding the mechanics behind the development of disease phenotypes, and for the screening of potential therapeutics. At Cellomatics, many of our scientists are trained molecular biologists, who have since specialised within one or more of our 5 therapeutic areas of research. To this end, they effectively utilise their extensive expertise to drive client projects across the fields of immunology, oncology, immuno-oncology, inflammation, and respiratory diseases through the application of both functional and analytical molecular tools.
An example of a functional molecular tool would be a cell line engineered to express a specific gene, to be used to assess phenotypic changes following exposure to a potential therapeutic. Meanwhile, an analytical molecular tool would be an endpoint assay such as qPCR or Western blot. The article aims to provide a flavour of the molecular capabilities available at Cellomatics.
Functional tools
At Cellomatics we employ molecular techniques and technology as tools to generate functional in vitro models of disfunction or disease. This primarily involves the specific manipulation of endogenous gene expression to produce transient, inducible, or stable cells displaying gene knockdown, integration, overexpression, and mutation. Dependent upon the client’s needs, such cell models can be utilised in proof-of-concept experiments to confirm pathway specificity and identify appropriate experimental readouts; to serve as physiologically relevant platforms for therapeutic mode of action investigations; to generate bespoke analytical assay platforms; or as positive or negative controls during therapeutic interrogation.
Plasmid Generation
Despite the advent of newer technologies like CRISPR-Cas9, traditional molecular cloning techniques are still vital for pre-clinical research. At Cellomatics we can engineer artificial plasmid DNA vectors to contain native, non-native, truncated, mutated, of tagged genes of interest. Through a range of techniques including transfection, transduction and Nucleofection we can successfully deliver these plasmids to both primary cells and immortalised cell lines. The resultant cell model can be used to study gene function in the context of assessing responses following stimulation with a compound of interest, the development of a disease phenotype, or target validation.
By way of brief explanation, plasmids are small circular DNA constructs that originate from bacteria, acquired through horizontal gene transfer, typically providing advantageous evolutionary gene expression such as antibiotic resistance. Molecular biology exploits this natural machinery to express exogenous genes in mammalian cells without the need for specific integration into the existing genome. With access to a large library of pre-generated gene expression plasmids and plasmid vector backbones, we can support both direct to cell experiments as well as plasmid synthesis, sub-cloning, and mutagenesis studies.
In the absence of a ready to transfect commercial plasmid, we can utilise ‘sub-cloning’ to create bespoke vectors. This can be essential to overcome size limitations of plasmid synthesis technologies, to add N or C-terminal epitope, fluorescent or biochemically responsive tags, or to introduce truncations or point mutations within a gene of interest sequence. To this end, the required building-block vectors are transformed and expanded in bacterial cultures, isolated using commercially available kits, and the sequences confirmed through Sanger sequencing or primer specific PCR amplification analysis. The desired plasmid components can then be combined through restriction endonuclease digestion, fragment isolation, clean up and ligation (Figure 1). A final round of bacterial transformation, expansion, isolation, and sequencing confirmation are performed before the plasmid vector is ready for use in experiments. For mutational work, reverse PCR and mismatched PCR amplification are a standard method.
Figure 1. Traditional molecular biology recombinant DNA techniques. Left to right: A miniprep of a plasmid vector. PCR amplification of 3 specific products of different molecular weight from 3 separate plasmid vectors. Restriction digests of plasmid DNA showing the release of products of different molecular weight, alongside the uncut plasmid vectors. A ligation of two restriction digest products, A and B, to generate a single chimeric construct (A+B). All images are of plasmid products run on ethidium bromide agarose gels of variable percentage. M=marker lane, P=plasmid, A/B/C/D/E = representative genes of interest
Transfection/Transduction methods
The visualisation of signalling pathways is a very common application of plasmid design and expression. Plasmids containing genes of interest fused to reporter genes such as luciferase or GFP can be used to monitor gene expression and subcellular localisation once introduced into cells.
The introduction of plasmids into cells is often accomplished by transfection, a term encompassing a range of techniques such as lipofection, calcium chloride transfection and electroporation. At Cellomatics, we have the capability to perform Nucleofection (a form of electroporation which transfers DNA directly into the cell nucleus), which has been shown to be capable of transfecting primary cells (e.g., immunological sub-populations of cells isolated from peripheral blood), suspension cells, and otherwise ‘hard to transfect’ cells.
Gene expression can also be achieved through viral transduction. Cellomatics utilises ready to transfect viral particles to deliver genes of interest to primary and immortalised cells.
One key advantage of including a visible tag in plasmid constructs is that gene expression can be easily detected by fluorescence microscopy following transfection. Moreover, the degree and localisation profile of gene expression can be monitored overtime and semi-quantified (Figure 2). Both primary and immortalised cell lines display differential susceptibility to various transfection methods, so it is essential to optimise this experimentally (Figure 2). Cellomatics has had success in employing both transient transfections and viral transductions to introduce DNA/RNA into cells for several applications including, but not limited to target validation, confirmation of therapeutic mode of action, generation of reporter cell lines, disease phenotype modelling, and functional assay controls/references.
Figure 2. Transient plasmid DNA transfection optimisation. A. Hela cells were transfected using lipofectamine reagent and GFPmax plasmid DNA complexes. Expression of GFP post-transfection was determined by both fluorescence intensity measurement and by fluorescence microscopy over a period of days. B. Cell lines MB231 and A549 were transfected with GFPMax plasmid DNA by Lipofection or Nucleofection. Approximately 24 hours are delivery of the plasmid, GFP expression was determined by way of fluorescence microscopy.
In contrast to the over-expression methods discussed above, Cellomatics also performs ‘knock-down’ gene expression for target validation and loss-of-function assays. Common techniques include siRNA and antisense RNA technology, and the initial transfection or transduction procedures are comparable to those described above. Successful gene knockdown can be analysed through the loss of a gene product either on a traditional agarose gel or via qPCR analysis, and through the loss of phenotypic activity mediated by the gene of interest (Figure 3).
Figure 3. siRNA transfection optimisation – qPCR analysis of target gene knockdown. A. DLD-1 and LoVo cells were transfected with 3 concentrations of a target gene siRNA, using DarmaFECT1 transfection reagent. B. DLD-1 and LoVo cells were transfected with 3 concentrations of a target gene siRNA, using RNAiMAX transfection reagent. Statistical significance was determined by way of comparison to the non-targeting control siRNA transfection performed with 30 µM siRNA (n=3, average +/-SEM).
Stable Expression methods
Transient transfections are useful for short-term studies; however, our team also has extensive experience generating polyclonal stable cell lines, via classical antibiotic selection, which express genes consistently over extended periods of time. The inclusion of a reporter gene and/or an antibiotic resistance sequence can be used to ‘select’ the sub-population of transfected cells expressing a gene of interest, and this forms the basis of generating stable cell lines. Typically, an antibiotic specific kill curve is first generated for each cell line identified for use in the stable cell line production. This process establishes the minimal concentration of antibiotic to kill all ‘untransformed’ cells over a given period, usually 7-14 days. Common methods to determine antibiotic killing are through viability assays and/or cell imaging (Figure 4). Conducting a kill curve ensures that during the stable cell line production, a sufficient selection pressure is applied to cells post-transfection so that only the sub-population expressing the desired construct remains.
Figure 4. Stable cell line production. A. An antibiotic kill curve performed over a period of 7 days, using 6 concentrations of antibiotic. Cell viability was determined by CellTiter-Glo® on day 2, 4, and 7 of selection. The percentage viability of each treatment condition was then calculated relative to the un-treated control condition for each given day. B. Live cell imaging of Wild-type and transfected HCT116 cells after 10 days of selection with Puromycin (2 µg/mL). Images captured in Brightfield at 4x magnification objective.
It is important to note that successful transfection does not guarantee expression of the desired gene. By nature, any polyclonal population will display a spectrum of expression. Dependent on the downstream experimental requirements, we can isolate and enrich stable monoclonal populations through serial dilution, cloning ring isolation, or FACS (flow cytometry assisted cell sorting) (Figure 5). The advantage of a monoclonal population is the uniformity of expression, growth and experimental response. However, their production can be time consuming and problematic as a large number of monoclonal populations must be profiled to find one with a suitable expression profile. Polyclonal and monoclonal cell lines can be validated by qPCR, western blot, flow cytometry, and by functional assays specific to the field of study (Figure 6).
Figure 5. Methods for stable cell line enrichment and monoclonal production. A. FACS purification and enrichment of double positive cells within a polyclonal population after antibiotic selection of a plasmid transfected cell line. Left to right: Wild-type cells remained negative for both the protein of interest and the associated fluorescent tag. The polyclonal population showed two distinct populations. Enrichment removed most of the negative and single positive cells. Purification was then performed on cells within the R5 ‘double positive’ quadrant, indicated by the red arrow. B. Serial dilution method to obtain stable monoclonal populations. Left to right: Flow cytometry analysis of Wild-type cells displayed no shift in the protein of interest compared to the isotype control. The polyclonal population displayed a mixture of cells expressing the protein of interest. The monoclonal population expanded from a single cell obtained by serial dilution of the polyclonal population displayed a clear increase in the expression of the protein of interest.
Figure 6. Confirmation of stable knockout cell line by western blot. Cell lysates obtained from 3 technical replicate cultures of a wild-type and shRNA knockdown population were analysed by western blot. Membranes were probed for expression of the target gene of interest and found to be knocked down in the population treated with shRNA and selected for 10 days.
Analytical tools
Gene expression may be altered in response to external or environmental stimuli, or through cellular processes leading to phenotypic changes associated with development or disease. This can be monitored at the DNA or RNA level; additionally, it is possible to assess changes in protein expression, function and interactions. These techniques can be used to determine the activation of cell signalling pathways. Samples can be derived from the functional platforms described above (e.g., for validation or to assess responses to stimulation), through cell models exposed to test compounds, or from primary or clinical material supplied by clients.
Analysis of RNA
The ability to multiplex readouts from targets within a small sample volume is a key factor in the successful application of the technologies we regularly employ when assessing gene expression changes in primary cells or clinical sample material derived from client studies.
Monitoring changes in gene expression, through the quantification of mRNA is a simple and moderately high-throughput method of molecular analysis. By way of multiplex qPCR or Quantigene analysis we can determine expression changes of 4-50 gene targets in a single assay. At Cellomatics Biosciences Ltd, we utilise published and experimentally established ‘biomarkers’ on these platforms to validate our in-vitro models of disease and inflammation. With well-defined and reproducible assay windows for our optimised biomarker genes, we can effectively assess the therapeutic activities of our clients’ test articles in these systems. This is particularly beneficial where sample quantity is limited, such as with primary immune cell assays, primary co-culture models, and patient derived material.
Quantitative real-time PCR (qPCR) functions through the conversion of mRNA to DNA, which is detected in a PCR reaction. The design of primers incorporating fluorescent probes and quenchers allows an increase in fluorescence related to the levels of specific PCR products to be visualised and quantified in real time, and assays for different gene products can be multiplexed and run together in the same sample. Data can then be expressed as fold-changes in gene expression, for example, in response to the stimulation of cells with a client’s proprietary compound (Figure 7). This technique can also be used to validate gene expression or knock-down following transfection.
Figure 7. qPCR analysis of Chondrocyte biomarker expression changes in response to the induction of Osteoarthritis. A. Real-time quantitative PCR amplification plots illustrating the detection of mRNA for GapDH and example genes of interest. B. Changes in Biomarker mRNA expression in a model of Osteoarthritis. Chondrocytes were stimulated with a cocktail containing IL-1β for 24 hours to induce the disease state of osteoarthritis. Five genes of interest were assessed for their suitability to serve as reliable biomarkers of the disease model. Graphs represent 3 technical replicates (n=3); mean ±S.E.M.
Alternatively, where sample volume is limited, gene expression is significantly low, and/or a large number of gene targets are to be analysed, a QuantiGene™ assay can be performed on either our MAGPIX or FLEXMAP 3D Luminex systems. QuantiGene™ assays function through the direct measurement of RNA using signal amplification rather than PCR-based target amplification. As part of Cellomatics work on Pulmonary fibrosis, Primary Human Lung Fibroblasts were analysed by QuantiGene™ to detect changes in extracellular matrix component genes after induction of fibrosis by TGFβ in the presence or absence of the therapeutic Pirfenidone (Figure 8).
Figure 8. Quantigene analysis of markers of fibrosis in Primary Human Lung Fibroblasts. Cell lysates were analysed using QuantiGene multiplex to detect changes in 4 extracellular matrix component genes, IHH, MMP-1, MMP-3 and MMP-13. Fold changes were determined by normalising target gene expression to house keeping genes (GAPDH and HPRT1). All treatment conditions were compared to the TGFβ treated samples (***p<0.001; **p<0.01; *p<0.05; n=3±SEM).
Analysis of protein expression
Gene expression changes form the foundations of how cells behave following exposure to potential new therapeutics, or how the disease phenotype develops. However, this is mediated by downstream changes in protein expression or post-translational modification. These changes can be monitored through techniques such as enzyme-linked immunosorbent assay (ELISA), western blot, or flow cytometry. These immunological techniques are based on a specific antibody binding to a protein of interest, which is linked to a range of detection techniques.
Western blotting can detect both changes in protein expression and post-translational modifications such as protein phosphorylation or proteolytic cleavage. For example, the expression of proteins such as Erk1/2 and subsequent changes in phosphorylation following cellular stimulation with EGF may be assessed. By using IRdye (infrared fluorescent dye)-labelled secondary antibodies, the detection of both the protein of interest and its phosphorylated form can be multiplexed (Figure 9).
Western blotting is also used as the endpoint of affinity assays, used to assess interactions between proteins which are either endogenously expressed or engineered into a cell line of interest. Here, a protein of interest is bound by a specific antibody which is then captured on a magnetic protein G Sepharose resin. Subsequent washing of the resin removes non-specific components of the lysate, effectively enriching for the bound protein of interest and any interacting proteins. After elution, the protein of interest and its associated proteins can be identified individually by western blot or collectively by LC-MS.
For example, the potential interaction between proteins ‘A’ and ‘B’ was investigated in wild-type and protein ‘A’ knockout A549 cells. Western blot analysis of whole cell lysate input samples confirmed the expression and absence of protein ‘A’ in wild-type and protein ‘A’ knockout cells respectively. Meanwhile, the expression of protein ‘B’ was confirmed in wild-type cells and to a much-reduced level in protein ‘A’ knockout cells (Figure 10). Co-IP using antibody against protein ‘A’ resulted in successful capture of protein ‘A’ and a small fraction of protein ‘B’. Co-IP using antibody against protein ‘B’ resulted in successful capture of protein ‘B’ but no detectable level of protein ‘A’. No immunoprecipitation was observed in knockout cells.
Figure 9. Detection and semi-quantitative analysis of phosphor-ERK by Western blot. Cells were stimulated for 5 minutes with 10 ng/mL EGF. Proteins were isolated and resolved by SDS-PAGE, followed by transfer to nitrocellulose membranes. Membranes probed for pErk 1/2 and Erk 1/2, and multiplexed detection was performed using IRdye secondary antibodies. Densitometric analysis was performed by normalisation of phosphor-ERK to the relative total ERK signal (mean of n=2 samples ±S.E.M., a t-test was used to determine statistical significance, where **P<0.01).
Figure 10. Co-IP to confirm protein-protein interaction in wild-type but not knockout cell line. Left panel: Whole cell lysate input western blot of protein ‘A’ and ‘B’ and housekeeping protein, GADPH. Right panel: Co-IP western blot of pull downs using no antibody, protein ‘A’ antibody, or protein ‘B’ antibody as indicated by the +/- key.
When accurate quantification and higher throughput are desired, ELISAs and Luminex assays are preferred to western blot. The Luminex system is suitable for the analysis of secreted chemokines and cytokines from a range of biological material, making it a staple assay we offer when conducting the assessment of immunology and inflammation therapeutics. ELISA’s can accurately quantify either internal or secreted proteins, as well as the quantity of certain post-translational modifications, such as protein phosphorylation, relative to total protein levels. For example, the expression profile of 4 targets was investigated in the presence or absence of TNF-α stimulation with Dexamethasone serving as an inhibitor of stimulation (Figure 11). Importantly, profiles were obtained at both the gene and protein level through QuantiGene™ and Luminex analysis respectively. The clear alignment of the stimulation and inhibition profiles accurately reflects the transcription-translation process of the central dogma.
Figure 11. Target gene and protein changes in response to TNF-α stimulation. A. QuantiGene™ analysis. Fold-change of target genes TNF-α, IL-6, IL-8, and MCP-1 expression normalised to the untreated control. B. Luminex cytokine analysis. Quantification (pg/mL) of target proteins, TNF-α, IL-6, IL-8, and MCP-1 expression.
Summary
The interaction of biological molecules forms the basis of how organisms are formed and function, and correspondingly they can be considered as the first point of investigation for how diseases develop and potentially druggable targets which may be utilised. Molecular biology allows us to exploit these phenomena by generating cell-based testing platforms, and by directly interrogating genetic responses in cells following stimulation with test compounds provided by clients.
