True ignorance is not the absence of knowledge, but the refusal to acquire it.
This article was prompted by a rather unpleasant exchange between myself and someone whose scientific knowledge is severely lacking. During this exchange, inevitably, the issue of genetic modification (GM) came up. When it did, I discovered that this individual, who is a sitting councillor for Limerick City North, opposes GM because he “doesn’t trust science under capitalism”. Such logic betrays not only a fundamental ignorance of what GM is, but also a conspiratorial mindset that is more befitting of Alex Jones and his acolytes..
Put simply, GM is a considerably more targeted version of artificial selection: we use it to make pest-resistant crops and life-saving recombinant protein drugs. Indeed, if I have type 1 diabetes, should I not take recombinant human insulin because we’re living under capitalism? The same applies to medicine more broadly: if I have a throat infection, should I not take antibiotics because we’re living under capitalism? What if I’m travelling to a country where polio is widespread: should I not get vaccinated because we don’t have a planned economy?
It should be stated, of course, that the profit motive is a major problem in R&D. Ben Goldacre and others have done an excellent job of exposing dubious practices in the pharmaceutical industry, for example. We should recognise, however, that this doesn’t mean the science itself is bad. We can fight for economic justice without alleging some vaguely evil conspiracy by geneticists, who have better things to be doing with their time. Indeed, if the reader wants some insight into the GM process, he can check out my final year project below. Alternatively, if the reader is short on time, GM can be summed up in seven easy steps:
1..We order a recombinant plasmid, i.e. a molecule of circular DNA into which our gene of interest has been inserted. For plant transformation, the most commonly used plasmid derives from the naturally occurring soil bacterium, Agrobacterium tumefaciens.
2..To confirm the identity of the plasmid, we induce its expression by growing it up in Luria Bertani broth. This broth contains ampicillin, meaning only plasmids designed with a resistance to ampicillin will survive.
3..Having purified our plasmid, we use a Miniprep Kit to break open the circular DNA. This is necessary to prepare the DNA for Polymerase Chain Reaction (PCR), which is our method of inducing the desired mutation.
4..In PCR, the double-stranded DNA is separated into two single strands. At this point, a pair of primers – short stretches of DNA – selectively bind to each strand. The forward primer, containing the altered nucleotide, binds to the bottom strand. The enzyme DNA Polymerase then synthesises new double-stranded DNA incorporating the mutation.
5..To confirm the success of mutagenesis, the new DNA is run on an agarose gel. It is then purified and ligated (sealed) to form a plasmid. Steps 2 to 5 are then repeated until all the desired mutations have been achieved.
6..The ligated plasmid is then mixed with plant cells or cut pieces of plants such as stems or leaves (explants). Some of the cells will take up the T-DNA, i.e. the section of the plasmid containing the mutated gene. The plasmid then inserts the T-DNA into one of the plant’s chromosomes, thus forming transgenic cells (GMOs).
7..The transgenic cells are then regenerated to form whole plants using tissue culture methods. The most viable samples are selected for the initiation of trial runs, which are performed in strict compliance with EPA regulations.
Using this method, we can modify corn to put up an effective resistance to Aspergillus, a mould that turns grains into health hazards. Through the induction of a few simple nucleotide changes, the corn cells are able to release specialised RNA molecules preventing the mould from releasing carcinogenic toxins. This is something which is only achievable through genetic engineering; there is only so much selective breeding can do. The same goes for recombinant protein drugs, which are produced by culturing E. coli cells transformed with plasmid mutants on a large-scale. Those who oppose GM ought to consider the plight of those with anaemia, who require recombinant erythropoietin (e.g. Epoetin alfa, manufactured by Amgen) to avoid chronic renal failure.
A vial of recombinant human insulin, used to treat diabetes by lowering levels of glucose in the blood.
‘Generation, Expression & Purification of Fluorescent Protein Variants’
Kareem Muhssin, B.Sc.
Submission Date: April 2014
Recombinant lectins are those whose glycan-binding properties have been altered by site-directed mutagenesis to recognise and selectively bind to specific glycan structures. By design, then, these lectins play an important role in the analysis of glycoprotein therapeutics, including the detection of glycosylation changes on the cell surface. The efficiency of these roles can be enhanced by fusing the lectins with fluorescent proteins such as EGFP and DsRed, for this enables the actual binding of glycans to be studied under fluorescence microscopy. By amplifying the fluorescence patterns of these fusion partners, we can optimise the visibility of labelled lectins; this is achieved by the induction of specific single-point mutations (SNPs). The induction of these SNPS, along with the induced expression of the resulting mutants and their purification by affinity chromatography, would comprise the bulk of the project.
In generating a palette of identical proteins with different emissions, we allow for more efficient labelling of lectins: for if the difference between the proteins is only a matter of two or three base-pairs, then any complications in the immobilisation of lectin-fluorescent protein fusions onto cell surfaces cannot be put down to differences in protein structure. In the author’s case, the parent protein was wild-type DsRed. The aim was to alter DsRed such that it emits green instead of red fluorescence; this protein is known as AG4. While the latter was not ultimately achieved, the author did obtain a much enhanced red mutant – known as E5up with V71M – which can now be used by the Irish Separation Science Cluster (ISSC) in their research. This research involves the detection of surface glycosylation changes – indicating apoptosis – in Chinese Hamster Ovary (CHO) cells under stress, and working to bring these cells back to a healthy state.
In producing a fluorescent protein with an enhanced emission pattern, the essential goal of the project was reached. In learning new scientific skills and refining existing ones, the broader goal was too.
List of Figures
Figure 1: The efficient immobilisation of an EGFP-lectin fusion onto the viscous surface area of a polyHIPE polymer
Figure 2: Flow diagram of the mutations used to obtain AG4 from DsRed
Figure 3: The GFP chromophore compared to its counterpart in DsRed
Figure 4: 1 kb DNA Ladder visualized by ethidium bromide staining on a 0.8% TAE agarose gel
Figure 5: Broad Range Protein Marker 2-212kDa visualised by Coomassie Blue staining on an SDS Polyacrylamide gel
Figure 6: Flow diagram of the physical work involved
Figure 7: Results of PCR confirmation gel for pQE-30, now carrying E5up
Figure 8: Results for transformation of concentrated JM109 with pQE-30 carrying E5up (enhanced red), viewed under the transilluminator
Figure 9: Contamination of GFP with DsRed (as expressed by JM109), viewed under the transilluminator
Figure 10: Results of PCR confirmation gel for pQE-30, presumed to be carrying E5
Figure 11: Results of 40-minute PCR confirmation gel for pQE-30, with twice the number of samples
Figure 12: Results of PCR confirmation gel for pQE-30, presumably now carrying E5
Figure 13: Results for transformation of JM109 with pQE-30 carrying E5 (enhanced red), viewed under the transilluminator
Figure 14: The results of agarose gel electrophoresis when the agarose hasn’t set properly
Figure 15: The amino acid sequence of DsRed-E5up, showing the replacement of Valine with Alanine at position 105
Figure 16: Cultures of JM109 expressing M3 Green (right) and M3 Red (left) in LBA, viewed under the transilluminator
Figure 17: Distorted SDS Polyacrylamide gel
Figure 18: M3 Red protein (presumed to be E5up, E5, or E57) after purification by immobilised nickel affinity chromatography
Figure 19: The results of SDS-PAGE for our M3 Red elutions, the filtered lysate, the unbound lysate, and the imidazole washes
Figure 20: Absorbance readings for our buffer-exchanged M3 Red elutions
Figure 21: Our M3 Red elution compared to presumed E57 under UV light
Figure 22: Results of a 0.7% agarose gel to confirm the presence of pQE-30 carrying M3 Red and pQE-30 carrying M3 Green
Figure 23: M3 Red (super-enhanced DsRed) compared to wild-type DsRed under the transilluminator
Figure 24: Fluorescence excitation and emission spectra of matured DsRed (~2 weeks at 25°C) compared to E5up
Figure 25: Absorbance readings for original DsRed elutions
Figure 26: Absorbance readings for our second series of M3 Red elutions
Figure 27: Results of SDS-PAGE for neat M3 Red and dilutions to 1 in 6
Figure 28: The amino acid sequence for M3 Red, demonstrating the induction of V71M and V105A, but not S197T
Figure 29: The amino acid sequence for our second mutant, showing the induction of V71M instead of S197T
Figure 30: Part of the amino acid sequence for M3 Green, demonstrating the induction of V71M and V105A, but not S197T
Figure 31: The amino acid sequence for M3 Red, with special emphasis on the residues preceding the Methionine at position 71
Figure 32: Part of the amino sequence for M3 Green, showing random mutations at positions 67-70
Figure 33: The complete amino acid sequence for M3 Green, divided across two reading frames
Figure 34: The results of SDS-PAGE for our M3 Red elutions
List of Tables
Table 1: The various mutants of DsRed, their phenotypic differences, and the mutations required to obtain them
Table 2: The three areas where DsRed needs improvement before it can be used for lectin fusions, compared to GFP
Table of Contents
Chapter 1: Introduction
- 1.1: The generation of fluorescent protein variants for use in the early detection of apoptosis
- 1.2: The immobilisation of lectin-fluorescent protein fusions onto cell surfaces
- 1.3: One example of the use of recombinant lectins in the analysis and purification of biotherapeutics
- 1.4: The various mutants of DsRed and the structure of the DsRed chromophore
- 1.5: Two short notes on site-directed mutagenesis
Chapter 2: Materials & Methods
- 2.1: Making up Luria Bertani (LB) broth / agar
- 2.2: Making up LB amp (LBA)
- 2.3: Preparing LBA agar plates
- 2.4: Preparing overnight cultures of KRX and JM109
- 2.5: Culturing plasmid DNA / E. coli expressing plasmid DNA
- 2.6: Plasmid DNA isolation
- 2.7: Agarose gel electrophoresis
- 2.9: Preparing glycerol stocks
- 2.10: Preparation of mutagenic primers for PCR
- 2.12: DNA purification from PCR mixtures
- 2.14: Making up Terrific Broth (TB)
- 2.15: Making up Super Optimal Broth (SOB)
- 2.16: Making up SOB with catabolite repression (SOC)
- 2.17: Running a SyBr Safe gel
- 2.18: DNA purification from agarose gels
- 2.20: Growing up competent cells of KRX and JM109
- 2.21: Transformation of competent cells
- 2.23: Sending plasmid DNA for sequencing
- 2.24: Preparing overnight cultures in TBA
- 2.25: Preparing an SDS Polyacrylamide gel
- 2.26: SDS-PAGE, staining & de-staining
- 2.27: Inducing protein expression
- 2.28: Making up lysis buffer
- 2.29: Stripping and recharging the IMAC resin
- 2.30: Cell lysis to obtain recombinant protein
- 2.32: Buffer exchange with PBS (protein purification)
Chapter 3: Diary of Work Done, Results & Data Interpretation
- 3.21 (Days 1–9): The induction of V105A and the expression of E5up
- 3.22 (Days 9–23): The induction of S197T and the expression of E5
- 3.23 (Days 23–28): The induction of V71M and the expression of AG4
- 3.24 (Days 28–35): Protein expression analysis & induced expression
- 3.25 (Days 35–47): Protein purification by nickel affinity chromatography
- Appendix A: The amino acid sequence of DsRed
- Appendix B: The nucleotide sequences of our mutagenic primers
- Appendix C: Table of the amino acids and their respective codons
- Appendix D: Diagram of the mutagenesis process and primer design
Chapter 1: Introduction
In the literature review, we concerned ourselves with the process of glycan evolution and how glycosylation can be used to enhance the efficacies of protein drugs. For the project, our focus was the use of site-directed mutagenesis to obtain enhanced variants of a wild-type fluorescent protein. These variants would be used by researchers at the ISSC in the early detection of apoptosis in eukaryotic cells. The onset of apoptosis is indicated by specific changes in glycosylation on the cell surface; thus, by fusing these fluorescent variants with lectins that bind to extracellular glycans, these changes can be monitored under fluorescence microscopy. The importance of glycosylation, therefore, was the ultimate bridge between the literature review and the project.
1.1: The generation of fluorescent protein variants for use in the early detection of apoptosis
Fluorescent proteins can be defined as structural homologues of the green fluorescent protein expressed by Aequorea victoria, a bioluminescent hydrozoan jellyfish, that share the ability to form a visible internal wavelength fluorophore from a sequence of three amino acids within their own polypeptide sequence. Fluorescent proteins are most commonly used for studying the dynamics and localisation of specific organelles (Rizzuto et al., 1995) or recombinant proteins (Ballestrem et al., 1998) in living cells. With regards to the latter, DsRed, a red fluorescent protein expressed by the Discosoma species of mushroom anemone, has proven especially useful.
The central aim of the project was thus to produce a series of DsRed variants with enhanced fluorescence patterns, by the successful induction of specific SNPs. We would achieve this by performing site-directed mutagenesis, inducing the expression of the resulting mutants and purifying them by Immobilised Metal Affinity Chromatography (IMAC). These fluorescent variants would then be used by researchers at the ISSC to create fusions with novel recombinant lectins, with the aim of detecting stress-induced glycosylation changes in CHO cells. In this endeavour, after the tagged cells have been treated with different stresses, flow cytometry is employed to detect any resultant changes in lectin binding.
In flow cytometry, lasers can be set for different wavelengths; therefore, the advantage of having mutants with different emissions is enormous. This is especially true if we want to label two glycoprotein drugs at the same time, or perform sequential labelling. The advantage is even greater when these mutants only differ by a few base-pairs, as would be the case with our DsRed mutants. Although our proteins would ultimately exhibit different fluorescences, the difference between them would only amount to two base-pairs, thus making them virtually the same. This allows for more effective labelling of lectins: for when obtaining data using flow cytometry, the possibility of interference or overlapping between different fluorescent proteins is always a major concern.
Apoptosis is a natural response in cells that are under stress, whether it is the stress of uncontrolled differentiation in cancer or the shear stress experienced by cells in a bioreactor. The onset of apoptosis is indicated by certain changes in glycosylation on the cell surface – for example, a decreased expression of O-glycosylated proteins (Batisse et al., 2004) and/or a marked decrease in cell surface sialylation (Shiratsuchi et al., 2002). Therefore, the ability to detect these changes early on is crucial in attempting to restore the cells to a healthy state (by altering the cell culture conditions). We can do exactly that by fusing recombinant lectins with fluorescent proteins and then immobilising these fusions onto the surfaces of cells to monitor any changes in lectin binding.
The importance of such an early detection method for the biopharmaceutical industry is obvious: for if we have a 10,000 litre vat of cells, and we want to determine if these cells are beginning to become stressed, then we need to so before the batch is compromised. At the moment, we have no choice but to take out a sample of cells, release the glycans, perform a mass-spec, and purify them by HPLC. It takes approximately 14 hours to carry this process out, by which time the cells are likely to be in trouble. There is no wonder, then, why so much focus is being put on developing a detection method that is fast and powerful – including the efforts of the ISSC.
1.2: The immobilisation of lectin-fluorescent protein fusions onto polymer and cell surfaces
Strictly speaking, fluorescent labelling is defined as the covalent attachment of a fluorophore to another molecule, such as a protein or nucleic acid. This is generally accomplished using a reactive derivative of the fluorophore that selectively binds to a functional group present on the surface of the target protein. Alternatively, recombinant DNA technology may be used to generate a fusion between the target protein and a naturally-fluorescent protein such as GFP or DsRed.
When we immobilise lectins, enzymes and other proteins onto the surfaces of cells, one of the easiest ways to check the efficiency of this immobilisation is via their fusion with fluorescent proteins. The likes of GFP and DsRed have the ideal amount and distribution of surface lysine residues for binding to a variety of chemically-activated surfaces. In addition, the fusion process itself is relatively simple to perform, involving the creation of a construct containing the fluorescent protein at either the NH2−terminus or the COOH-terminus of the protein of interest and the insertion of this construct into an appropriate expression vector, which is accomplished by ligation into a multi-cloning site. To quote Snapp et al. (2005) on the final design of the construct:
The final FFP [fused fluorescent protein] construct will contain an in-frame fusion between the FP and the protein of interest, an unambiguous initiating methionine within the appropriate Kozak sequence (for eukaryotes, this is 5′-ACCATGG-3′, where the internal ATG is the initiating methionine), a linker between the FP and the protein of interest (if necessary), and appropriate regulatory elements.
There are three bonuses to fusing lectins with fluorescent proteins:
1. By virtue of being visible, these proteins can give us a direct measurement of the efficiency of lectin immobilisation. This is determined quantitatively by the percentage of the fluorescent protein that has bound to the surface, and if this protein is evenly distributed. Figure 1 displays the immobilisation of an EGFP-lectin fusion onto the surface of a PolyHIPE polymer under 10X microscopy. PolyHIPEs are porous polymers with highly viscous surface areas that enable their use in reaction supports, separation membranes and other applications.
Figure 1: The efficient immobilisation of an EGFP-lectin fusion onto the viscous surface area of a PolyHIPE polymer. Image sourced from Audouin et al. (2012): ‘Polypeptide grafted macroporous PolyHIPE by surface initiated N-Carboxyanhydride Polymerization as a platform for bioconjugation’. Macromolecules, 45, 6127-6135.
In the diagram, we can see a nice distribution of the fluorescent protein, indicating a high efficiency of lectin immobilisation onto the polymer surface. This efficiency tells us that we don’t have to label each lectin in a different way: rather, we now have a generic way to do so.
2. Fluorescent proteins have often been found to stabilise certain lectins.
3. Orientation-specific immobilisation becomes more possible, as when we immobilise a fluorescent protein partner onto a particular lectin, we are making sure that the lectin binding pocket is now orientated correctly. (The protein is always fused to an N- or C-terminus, so it doesn’t interfere with the binding pocket.)
1.3: One example of the use of recombinant lectins in the analysis and purification of biotherapeutics
In the production of biopharmaceuticals, the maximum possible level of purity is desirable. By labelling our protein of interest with an affinity tag – be it a short peptide sequence or a full protein – we can precipitate the final product from the thousands of contaminants involved with relative ease. For the analysis of glycoprotein therapeutics, labelling with recombinant lectins – sometimes fused to fluorescent proteins – is considered the most suitable option. Indeed, the novel recombinant lectins developed by the ISSC are much better than commercial lectins at binding to specific targeted therapeutics.
One example of a recombinant lectin that has proven useful in the analysis and purification of biotherapeutics would be the PA-IL lectin, expressed by Pseudomonas aeruginosa. As part of a recent collaborative study by the ISSC and Dublin City University (DCU), specific amino acid residues located in the carbohydrate binding site of a recombinant PA-IL protein (rPA-ILNmE6) were substituted at random by site-directed mutagenesis (Keogh et al., 2014). This resulted in high affinity and specificity of the protein for N-acetyl-lactosamine (LacNAc) and for β-linked galactose exhibited by N-linked glycans on the surfaces of glycoproteins (Keogh et al., 2014).
The researchers demonstrated the use of this novel recombinant prokaryotic lectin in the analysis of glycoproteins and in the isolation and selective fractionation (purification) of glycoproteins and their glycoforms. One way of doing this involved immobilisation of the lectin onto magnetic particles (Keogh et al., 2014). To quote from the paper:
To evaluate the ability of these lectin functionalized magnetic particles to isolate glycoproteins displaying terminal β1–4 galactose, pull down assays were performed in 1.5mL tubes, using a test protein mixture prepared by mixing asialotransferrin with recombinant green fluorescent protein (GFP) and RNase B. Fractions of unbound and bound protein were ultimately evaluated by SDS-PAGE. [From this gel], it could be clearly seen that the rPA-ILNmE6 magnetic beads selectively and efficiently extracted the asialotransferrin from the protein mixture.
1.4: The various mutants of DsRed and the structure of the DsRed chromophore
DsRed was first discovered in late 1999 by a research team of the Russian Academy of Science, who observed that reef Anthozoa express fluorescent proteins with hues ranging from red to cyan (Matz et al., 1999). Soon afterwards, wild-type DsRed was found to have certain properties that prevent its efficient use as an in vivo reporter. These drawbacks, along with obtaining a palette of DsRed mutants to work with, are what necessitate site-directed mutagenesis. They include the following:
1. When compared to wild-type GFP, Baird et al. (2000) and Terskikh et al. (2001) observed a slow rate of fluorescence maturation for DsRed. This difference is most striking at lower temperatures (just below room temperature) – temperatures at which GFP has maximum folding efficiency (Kimata et al., 1997). This limits the utility of DsRed for studies of heterologous expression systems that are maintained at low temperature (such as Drosophila or Xenopus).
In contrast, DsRed and all other Anthozoa proteins studied thus far mature faster than GFP at temperatures higher than 30°C (Terskikh et al., 2001). These temperature-dependent rates could be evolutionary adaptations to the animals’ natural habitat: corals live in the warm Indo-Pacific waters, while Aequorea victoria lives in the cold boreal waters (Terskikh et al., 2001).
2. Even after lengthy periods of maturation (several weeks), wild-type DsRed retains residual green fluorescence at 500 nm, amounting to 4–5% of the major red fluorescence peak (Terskikh et al., 2001). This can be a major source of problems in multicolour detection applications.
Both of these issues can be overcome by the successful induction of specific SNPs. By the induction of V105A in wild-type DsRed, we can accelerate fluorophore maturation such that not only is the red fluorescence enhanced, but any residual green fluorescence is eliminated entirely. Presumably, substituting a bulky Valine for a more compact Alanine causes the overall structure to relax, facilitating the complete maturation of the red fluorophore (Terskikh et al., 2001). The resulting mutant is known as E5up.
It is also possible to obtain DsRed mutants exhibiting intense green fluorescence, if so desired. One such mutant, AG4, can be achieved by the induction of V71M in E5 (i.e. DsRed carrying both V105A and S197T). The induction of S197T enhances green fluorescence; the induction of V71M abrogates the enhanced red of V105A, thus enabling the DsRed mutant to fluoresce green. Most likely, the introduction of a larger side chain that comes with substituting Methionine for Valine produces substantial distortion of the fluorophore environment, preventing complete maturation of the red fluorescence (Terskikh et al., 2001).
Figure 2 represents how AG4 can be obtained from DsRed by the successive induction of V105A, S197T and V71M:
Figure 2: Flow diagram of the mutations used to obtain AG4 from DsRed.
The author of this report opted to obtain AG4: for while the production of yet another green fluorescent protein may not seem to be of any advantage initially, in reality, it can be used as an ideal counterpart for DsRed in the multi-coloured labelling of novel recombinant lectins. The minimal sequence differences between AG4 and DsRed ensure their similar behaviour on cell surfaces, i.e. precipitation, non-specific accumulation, and interaction with cellular components (Terskikh et al., 2001). Thus, any observable differences in the behaviour of two proteins – one labelled with DsRed, the other with AG4 – should be attributed to the proteins themselves, and not the fluorescent tags. Featured below is a table of the DsRed mutants and their phenotypic properties:
Table 1: The various mutants of DsRed, their phenotypic differences, and the mutations required to obtain them.
It is important that we elaborate somewhat on the structure of the DsRed chromophore, for it is by site-directed mutagenesis in and around the chromophore that the red fluorescence is altered. Fluorescent proteins with different emission patterns amount to proteins with different chromophore environments.
The DsRed chromophore is an auto-catalytically produced variant of the GFP chromophore. As shown in Figure 3, the main difference between the two is that the DsRed chromophore has an additional double bond (drawn in yellow), which extends its conjugation and causes the shift to red fluorescence. Robert E. Campbell of the University of Alberta provides a more detailed explanation in this regard (2007):
The key difference between the red and green proteins is that Discosoma red fluorescent protein undergoes an additional fourth step in the chromophore maturation pathway, oxidizing the adjacent Cα-N bond to form an acylimine moiety that extends the conjugated system by two double bonds.
Figure 3: One double bond – the result of pi-electron conjugation – is primarily what distinguishes the DsRed chromophore from that of GFP. Featured here are simplified versions of the structures of O. Shimomura (1979): ‘Structure of the chromophore of Aequorea green fluorescent protein.’ FEBS Letters 104.
In October 2000, Gross et al. of the University of Pennsylvania investigated the structure of the DsRed chromophore. From this study, they were able to identify key differences between the major properties of wild-type DsRed and wild-type GFP. Of these properties, the authors singled out three areas where DsRed needs improvement before it can be used as an efficient fusion partner: namely, suppressing its aggregative tendencies, and increasing both the completeness and the speed of maturation to the desired red form (Gross et al., 2000). These are summarised in the table below:
Table 2: The three areas where DsRed needs improvement before it can be used for lectin fusions, compared to GFP. Abridged version of the table drawn up by Gross et al. (2000): ‘The structure of the chromophore within DsRed, a red fluorescent protein from coral.’ PNAS, vol. 97 no. 22, 11990–11995.
With regards to the extent and speed of chromophore maturation, although the residual green fluorescence in wild-type DsRed does not show up in the emission spectrum – due to fluorescence resonance energy transfer (FRET) within the tetramer – its excitation peak will nonetheless contribute spectral cross-talk in experiments that use GFP as a separate label or FRET donor (Gross et al., 2000). Therefore, if DsRed is to be used as a fluorescent label, then increasing the speed and the extent of chromophore maturation – thereby eliminating residual green fluorescence – is of crucial importance. By the induction of V105A, this is what we had hoped to achieve in the project.
1.5: Two short notes on site-directed mutagenesis
Site-directed mutagenesis is a convenient method for inducing a specific mutation at a specific site in a sequence of plasmid DNA. This mutation can be a substitution, deletion or insertion. Site-directed mutagenesis has many uses, such as the removal of restriction sites from a plasmid, investigating the function of specific amino acids in an enzyme, or amplifying the phenotypic properties of a particular protein.
With regards to PCR-based site directed mutagenesis, the principle is to design a pair of PCR primers adjacent to one other, such that it is the entire plasmid that is amplified by PCR. One of these primers – the forward primer – incorporates the desired mutation. The result of the PCR is a linear product whose ends can then be sealed together (after phosphorylation) with DNA ligase. The ligated vector is then taken up and expressed by E. coli (transformation). This process can be summarised in seven steps:
Step 1: Primer Design
Step 2: PCR
Step 3: Purification of the PCR product
Step 4: Phosphorylation of the 5’ termini (if necessary)
Step 5: Ligation of the DNA ends
Step 6: Transformation of competent E. coli
Step 7: Confirmation of success by DNA sequencing.
Short notes on primer design are included in Section 3.1 of this report. A detailed diagram on primer design and the mutagenesis process is also included in Appendix D.
Chapter 2: Materials & Methods
2.1: Making up Luria Bertani (LB) broth / agar
Prepare the required volume of distilled water in a beaker. Place the beaker on a heat-stir, with a magnet bar resting at the bottom. Set the heat-stir to the second-lowest setting. Weigh out the appropriate amounts of Tryptone, NaCl and Yeast Extract and add them to the beaker. (For making any volume of LB, these components are added at concentrations of 10 g/L.) Once the solution is homogenous, pour it into a graduated cylinder and top up to the correct volume with distilled water. Pour this into a duran that has been clearly labelled and sealed with autoclave tape. If making LB agar, it is at this point that agarose is added (Scharlau Agar Bacteriological 07-004) at a concentration of 15 g/L. The agarose is allowed to settle at the bottom of the jar.
Autoclave the jar, keeping the lid loose to ensure the LB is heated properly. (Check the volume of water in the autoclave before proceeding. Once the lid is shut, switch on the device and set the knob to steam.) The heating process takes approximately 30 minutes. Once the pressure has fallen below half a bar, set the knob to ‘exhaust’ and wait for the pressure to hit zero. At this point, open the lid and retrieve the LB. Once the autoclaved LB has cooled down (usually after 30-40 minutes), it is ready for use.
2.2: Making up LB amp (LBA)
Prepare a 100 mg/mL concentration of ampicillin in a sterilin tube (e.g. 0.359 g ampicillin in 3.59 mL distilled water). Aliquot 500 μL volumes of this preparation into a series of microfuge tubes. To make LB amp, add ampicillin to LB at a concentration of 1 μL/mL. (It is good scientific practice to do this in the fume hood.) The LBA is now ready for use.
2.3: Preparing LBA agar plates
In the fume hood, prepare a series of empty plates in close proximity to the Bunsen flame. Slowly and gently pour the LBA agar into each plate, making sure to cover each plate’s surface area. Keep the plates by the flame until they have settled, with the lids partially covering each plate. Close the plates once the agar has settled. The plates are now ready for use.
2.4: Preparing overnight cultures of KRX and JM109
Culture KRX and JM109 (ordered from PROMEGA) by inoculating 10 μL of each sample in two sterilin tubes of 5 mL LB. Prepare another tube of sterile 5 mL LB as a control. (It is good scientific practice to do all of this in the fume hood.) Store the tubes overnight at 37˚C in an incubator shaker. The following day, retrieve the tubes and examine their appearance. Successfully inoculated samples appear murky in colour. A cloudy appearance for LB (or LBA) that has not been inoculated indicates contamination.
2.5: Culturing plasmid DNA / E. coli expressing plasmid DNA
Add 10 μL of plasmid (or a single colony expressing the plasmid) to 5 mL LBA in a sterilin tube. Incubate overnight at 37˚C in an incubator shaker. The following day, retrieve the tubes and examine their appearance.
2.6: Plasmid DNA isolation
Sigma GenElute Plasmid Miniprep Kit
Transfer 1.5 mL of cell culture (expressing the plasmid) to a microfuge tube. Centrifuge the tube at 13,000 rpm for 2 minutes to pellet the cells. Remove the supernatant and resuspend the pellet in 200 μL of resuspension solution (kept in the fridge, as it contains RNase A solution). Add 200 μL of lysis solution. Gently invert the tube and leave to incubate at room temperature for 5 minutes. (Harsh mixing may cause the co-purification of unwanted chromosomal DNA.) Add 350 μL of neutralisation solution and mix by inversion to precipitate cell debris, lipids, proteins and chromosomal DNA. Leave the mixture at room temperature for 10 minutes. Collect the precipitate by centrifugation at 13,000 rpm for 10 minutes.
Prepare a spin column for binding by adding 500 μL of column preparation solution and centrifuging at 13,000 rpm for 1 minute. Transfer the supernatant to the prepared spin column and centrifuge for 30 seconds to bind the plasmid DNA. Discard the flow-through and add 750 μL of wash solution. Centrifuge at 13,000 rpm for 30 seconds to remove any contaminants. Discard the flow-through and dry the matrix by centrifuging at 13,000 rpm for 2 minutes. Transfer the spin column to a new microfuge tube and add 100 μL of distilled water. Elute the plasmid DNA by centrifugation at 13,000 rpm for 30 seconds.
2.7: Agarose gel electrophoresis
Prepare a 0.7% agarose solution by adding 0.7% agarose to TAE buffer and dissolving by boiling. The agarose is stored at 60˚C to prevent solidification. When required, the heated agarose is poured into plastic trays and allowed to set containing a plastic comb to form wells. 1X TAE buffer is used as the running buffer. Typically, 2 μL of gel loading dye is mixed with 7 μL of sample to help the sample settle in the well. A 1kb DNA ladder is used:
Figure 4: 1 kb DNA Ladder visualized by ethidium bromide staining on a 0.8% TAE agarose gel. © New England Biolabs Inc.
The ladder is also loaded with gel loading dye. Gels are run at 120 volts for 20-40 minutes. The gel is then stained for 15 minutes by immersion in an ethidium bromide solution (wearing gloves, as ethidium bromide is carcinogenic). Gels are visualised using the UV transilluminator. Dispose of gels and gloves in the nearest ethidium bromide waste bin.
2.8: Making up TE Buffer
TE Buffer consists of a 10mM concentration of Tris-HCl and a 1mM concentration of Na2EDTA in 100 mL of distilled water. Calculate the amount in grams needed for each concentration, weigh them out, and add them to a duran of 100 mL distilled water. Using a dropper, adjust the pH to 8.0 with HCl. The TE Buffer is now ready for use. Example:
Tris-HCl has a molecular weight of 121.14 g/M. Therefore, for a 10mM concentration, weigh out 1.21 g. The working assumption here is a 1L volume; thus, for 100 mL, divide by 10 once more to get 0.121 g.
2.9: Preparing glycerol stocks
In the fume hood, prepare cryogenic vials in duplicate, each containing 0.5 mL of glycerol and 1 mL of competent cells in LBA / plasmid in LBA. Store the vials at -80˚C.
2.10: Preparation of mutagenic primers for PCR
The forward and reverse primers for each mutation are resuspended in the appropriate volumes of TE Buffer (amount of oligo X 10). Prepare 1 in 10 dilutions of these stocks (10 μL stock + 90 μL TE Buffer) in clearly labelled microfuge tubes. Use the stickers provided with the information sheet for each primer to label the tubes.
2.11: Mutagenesis PCR
PCR reactions are performed in the Veriti 96 well Thermal Cycler (Applied Biosciences). The reaction components are as follows:
Template DNA 1 μL
dNTPs (10 μM) 1 μL
Primers (10 μM) 1 μL of each
Buffer (5X) 10 μL Q5 High Fidelity
Distilled water 35 μL
DNA polymerase 1 μL Q5 High Fidelity
Prepare a master mix containing 4X each component for the four potential annealing temperatures (55, 60, 65, 70˚C). (The dNTP mix contains 5 μL of each dNTP in 30 μL of distilled water. The DNA polymerase is added last.) Aliquot 50 μL of the master mix into 4 PCR tubes each. Pulse the tubes to ensure the reaction is concentrated at the bottom. Program the Veriti 96 well Thermal Cycler as follows:
Browse/New methods → 2 minute extension → View/edit → Zones → Change to 3 minute extension → Run (check 50 μL) → Start PCR.
The 5X Buffer and DNA polymerase are stored at -20˚C.
2.12: DNA purification from PCR mixtures
After running a confirmation gel (5 μL for each PCR reaction), there should be 45 μL remaining in each PCR tube. Combine the samples that appear on the gel for 90 μL in one microfuge tube. Add 500 μL of capture buffer type 3 and mix the sample thoroughly by vortexing. Incubate the tube at 55˚C for 10-15 minutes until the agarose gel is fully dissolved. During this incubation, the tube is inverted every 2-3 minutes. Place the sample in a spin column and centrifuge at 13,000 rpm for 30 seconds. Discard the flow-through. Add 500 μL of wash buffer type 1 and centrifuge at 13,000 rpm for 30 seconds. Discard the flow through and dry the column in a new microfuge tube by centrifugation at 13,000 rpm for 2 minutes. Add 30 μL of distilled water to the centre of the column matrix and leave at room temperature for 2 minutes. Centrifuge the column at 13,000 rpm for 30 seconds to elute the DNA. The purified DNA is then stored at -20˚C.
2.13: Restriction digest
For a total of volume of 50 μL, prepare each reaction as follows:
Buffer (10X) 5 μL Buffer 4
Purified plasmid 30 μL Linear PCR product
Enzyme (DpnI) 1.5 μL 20,000 units/mL
Distilled water 13.5 μL
The restriction enzyme is added last. The reactions are incubated at 37˚C for 2 hours.
2.14: Making up Terrific Broth (TB)
For a 1L volume of TB, prepare the following in 900 mL of distilled water:
Tryptone 12 g (Scharlau Casein Trypsic Peptone 07-119)
Yeast extract 24 g (Scharlau Yeast Extract 07-079)
Glycerol (50%) 4 mL
Autoclave the prepared mixture. After cooling, add 100 mL of 1M potassium phosphate buffer aseptically. The TB is now ready for use. (Some of the phosphate buffer may aggregate to form crystals. It is easy to mistake this as contamination.)
2.15: Making up Super Optimal Broth (SOB)
Prepare the following in the appropriate amount of distilled water:
Tryptone 20 g/L (Scharlau Casein Trypsic Peptone 07-119)
Yeast extract 5 g/L (Scharlau Yeast Extract 07-079)
NaCl 500 mg/L
KCl 2.5 mM
The molecular weight of KCl is 74.55 g/M; therefore, for a 0.0025M concentration, we need 0.1864g. The working assumption here is 1L, so for 500 mL of SOB (for example), divide by 2 to get 0.0932g. Add this amount to 500 mL of distilled water and autoclave.
Once the SOB has cooled, we need to add 2 mL of MgCl2 and MgSO4, at concentrations of 1M. (The molecular weight of MgCl2 is 95.22 g/M. The working assumption is 1L, so if we make up 20 mL, for example, we divide by 50 to get 1.9044g. Do the same for MgSO4.) Once these have been added, the SOB is ready for use.
2.16: Making up SOB with catabolite repression (SOC)
In the fume-hood, add 1 mL of filter-sterilised MgCl2 and MgSO4 to 100 mL of SOB. Proceed to add 2 mL of filter-sterilised glucose (50%). The SOC is now ready for use.
2.17: Running a SyBr Safe gel
Prepare a 0.7% agarose gel as normal. 3 μL of SyBr Safe is added to the gel before it sets. Add 8 μL of loading dye to each 50 μL digest. Once the gel has set, remove the comb and load 25 μL each of these samples. Run the gel at 120 volts for 20-40 minutes. View the gel under the transilluminator to identify the best expressed plasmid.
2.18: DNA purification from agarose gels
The DNA band to be purified from the SyBr Safe gel is excised using a scalpel. 500 μL of capture buffer type 3 is added and the sample is mixed thoroughly by vortexing. The tube is incubated at 55˚C for 10-15 minutes until the agarose gel is fully dissolved. During this incubation, the tube is inverted every 2-3 minutes. The sample is placed in a spin column and centrifuged at 13,000 rpm for 30 seconds. The flow-through is discarded. 500 μL of wash buffer type 1 is added and centrifuged at 13,000 rpm for 30 seconds. The flow-through is discarded, and the column is dried in a new microfuge tube by centrifugation at 13,000 rpm for 2 minutes. 30 μL of distilled water is added to the centre of the column matrix and left at room temperature for 2 minutes. The column is then centrifuged at 13,000 rpm for 30 seconds to elute the DNA. The purified DNA is then stored at -20˚C.
For a total of volume of 50 μL, prepare each reaction as follows:
Purified plasmid 40 μL
Buffer (10X) 5 μL T4 DNA ligase buffer
Ligase (T4) 2 μL T4 DNA ligase
Distilled water 3 μL
Pulse each tube, and incubate at room temperature for 3 hours.
2.20: Growing up competent cells of KRX and JM109
Add 2 mL of KRX and JM109 overnight cultures to 200 mL of SOB (to which 2 mL of MgCl2 and MgSO4 have been added). Incubate for roughly 3 hours in an incubator shaker. Measure the absorbance at 1.5 hour intervals, with the spectrophotometer set to 600 nm. Once the absorbance has reached 0.4 – 0.6, pour each culture into sterile centrifuge bottles and centrifuge at 4,500 rpm for 5 minutes to pellet the cells. Decant the supernatant and resuspend the cells in 80 mL of cooled TB buffer. Leave the resuspended solution on ice for 10 minutes. Pellet the cells by centrifugation at 5,500 rpm for 5 minutes. Decant the supernatant, and gently resuspend the cells in 15 mL of chilled TB buffer. Slowly add 7% DMSO, incubate the suspension on ice for 10 minutes. Prepare aliquots of 400 μL in sterile 1.5 mL microfuge tubes, and flash freeze them using a block that has previously been cooled to -80˚C. The competent cell aliquots, now ready for use, are then stored at -80˚C.
2.21: Transformation of competent cells
The aliquots of competent cells are thawed on ice. 200 μL of competent cells are gently mixed with 5 μL of plasmid DNA in a sterile microfuge tube (fume-hood). The tube is incubated on ice for 30 minutes to allow the DNA and cells to bind. The cells are heat-shocked at 42˚C for 30 seconds and put back on ice for 2 minutes. 800 μL of SOC broth is added to the cells (fume-hood) and they are incubated at 37˚C for 1 hour. 200 μL of this neat suspension is pipetted on to an LBA agar plate and is incubated overnight at 37˚C. The remaining 800 μL is centrifuged at 5,000 rpm for 60 seconds. 600 μL of supernatant is removed; the pellet is resuspended in the remaining 200 μL and spread onto an LBA agar plate, to be incubated overnight at 37˚C. A control plate is also prepared using 200 μL of competent cells that haven’t taken up the plasmid.
2.22: Streak plates
Sterilize the inoculation loop by passing it through a flame. When the loop has cooled, use it to pick a single colony of E. coli. Drag the inoculation loop across the edges of the agar surface until roughly 30% of the plate has been covered. Re-sterilize the loop each time, and rotate the plate 90 degrees. Drag the loop in a zig-zag pattern from the final streaked area towards the starting point, but do not connect the two. Incubate the plates overnight at 37˚C.
2.23: Sending plasmid DNA for sequencing
Aliquot 15 μL of purified plasmid DNA into two microfuge tubes each. Having paid for sequencing services online, label each tube correctly and send to the following address:
Anzinger Str. 7a,
Upon receipt of the sequencing results, EXPASY is used to obtain the amino acid sequences, and thus to determine the success of site-directed mutagenesis.
2.24: Preparing overnight cultures in TBA
8 μL of amp is added to 8 mL of TB in a sterilin tube. 5 μL of IPTG is then added. The TBA is then inoculated with a looped colony of competent cells. Prepare another tube containing 8 mL of sterile TBA as a control. (It is good scientific practice to do all of this in the fume hood.) Store the tubes overnight at 37˚C in an incubator shaker. The following day, retrieve the tubes and examine their appearance. Successfully inoculated samples appear murky in colour. A cloudy appearance for TBA that has not been inoculated indicates contamination.
2.25: Preparing an SDS Polyacrylamide gel
Prepare two SDS-PAGE plates. Place a rubber gasket ridge-side-up around the furrowed plate. Lay the smooth plate on top of the latter, and secure them both with a clip on each side. Use distilled water to test if the plates are air-tight. Prepare a 15% resolving gel and a 4% stacking gel as follows:
15% resolving gel 4% stacking gel
30% Acrylamide (mL) 3.75 0.325
Distilled water (mL) 1.75875 1.54
1.5M Tris-HCl pH 8.8 (mL) 1.875 –
0.5M Tris-HCl pH 6.8 (mL) – 0.625
10% APS (μL) 37.5 12.5
10% SDS (μL) 75 25
Add 3.5 μL of Temed to the resolving gel and pour it in. Add 0.5 – 1 mL of 20% IMS on top to resolve any bubbles, and wait approximately 40 minutes for the gel to set. At this point, add Temed to the stacking gel, pour out the IMS, and pour in the stacking gel. Slowly and gently slot in the comb, and wait roughly 30 minutes for the gel to set. If the gel is not to be used immediately, cover it with some paper towels, soak it with distilled water, and store it in a sealed bag at 4˚C.
2.26: SDS-PAGE, staining & de-staining
Prepare the SDS-PAGE box by connecting it to a power station and pouring in 1X SDS-PAGE buffer to the wire. Slot in the prepared gel at the front, with a flat plate at the back. Secure the plates, fill the box to the top with 1X buffer, and remove the combs. Run at 15 mA for 30 minutes. Prepare 1 in 5 dilutions (0.8 mL distilled water + 0.2 mL sample) and read the absorbance at 600 nm for each. Use this figure to calculate the volume of culture to be harvested as follows:
(0.7 / Absorbance X 5) X 300 = Volume of culture (μL)
Aliquot these volumes into new microfuge tubes, and top them all up to the highest value with distilled water. Centrifuge these samples at 13,300 rpm for 2 minutes. Discard the supernatant, and resuspend the pellet in 50 μL of 10X SDS buffer.* (Do this in the fumehood, while wearing gloves.) Heat the samples at 100˚C for 5 minutes.
Load 18 μL in each lane, as well as the ladder in at least one lane, and run the gel at 70 V for 10-15 minutes. A 2-212kDa protein ladder is used:
Figure 5: Broad Range Protein Marker 2-212kDa visualised by Coomassie Blue staining on an SDS Polyacrylamide gel. © New England Biolabs Inc.
Once the samples have run out, ramp up the voltage to 120 V. Once the dye has run off – usually after 90 minutes – gently transfer the gel to an empty butter box. Stain the gel using Coomassie Blue staining solution.
To de-stain the gel, pour out the Coomassie Blue solution (reusable) and allow the gel to be washed in distilled water for 15 minutes. Discard the water and pour in Coomassie de-staining solution, leaving the gel on the shaker for the same amount of time it was left to stain. Pour out the de-staining solution and examine the gel.
* The procedure for protein fractions is the same, except that instead of pelleted cells being resuspended in 50 μL of 10X SDS buffer, 2 μL of 10X SDS buffer is added to each 18 μL of protein sample to be loaded.
2.27: Inducing protein expression
2 mL of the clone identified during screening is used to inoculate 200 mL of TBA. This culture is incubated in an incubator shaker at 37˚C, with the absorbance (600nm) read every hour. When this reaches 0.4 – 0.6, 200 μL of IPTG is added aseptically and the culture is transferred to a 30˚C incubator shaker for 16 hours.
2.28: Making up lysis buffer
NaH2PO4 50 mM
NaCl 0.5 M
Imidazole 20 – 250 mM
2.29: Stripping and recharging the IMAC resin
Note: 1 Column Volume (CV) = 2 mL.
The column is first washed with 2 CV of distilled water, followed by 2 CV of 50% ethanol. The metal ions are then stripped by washing with 2 CV of 100 mM EDTA, pH 8.0. To remove any remaining impurities, the column is then washed with 2 CV of 200 mM NaCl, 2 CV of distilled water and 10 CV of 30% isopropanol. The resin is then washed with 10 CV of distilled water and recharged with half a CV of 100 mM NiSO4. The column is again washed with 10 CV of distilled water and stored in 20% ethanol.
2.30: Cell lysis to obtain recombinant protein
Having induced expression with IPTG, the culture is retrieved and centrifuged at 5,000 rpm for 10 minutes at 4˚C using sterile centrifuge bottles. The supernatant is poured off and the pellet is resuspended in 100 mL of lysis buffer (20 mM imidazole) with 200 μL of anti-foam. The cells are lysed using a cell disruptor (Constant Systems Ltd.) at 15 kPSi twice. This cell lysate is collected in high speed centrifuge tubes and spun at 13,000 rpm for 40 minutes at 4˚C. The lysate is filtered through a Whatman filter into a clean duran.
Step Added Fraction Collected
- Rinse 10 CV Distilled water –
- Equilibrate 10 CV Lysis buffer (20 mM) –
- Lysate Filtered lysate Unbound lysate
- Wash 10 CV Lysis buffer (20 mM) 20 mM wash X 100 mL
- Wash (alt.) 10 CV Lysis buffer (80 mM) 80 mM wash X 100 mL
- Elution 5 CV Lysis buffer (500 mM) 12 X 1 mL
- Rinse 10 CV Distilled water –
- Storage In 20% IMS –
The protein fractions are visibly inspected using a transilluminator, to confirm their emission patterns.
2.32: Buffer exchange with PBS (protein purification)
Having inspected the protein elutions under UV light, the most concentrated fractions are buffer exchanged into 1X PBS for further spectral and visual analysis. Do this as follows:
Aliquot 500 μL of protein elution into a Vivaspin column, and centrifuge at 13,000 rpm for 10 minutes. Repeat this until all of the protein has collected in the filter membrane (10,000 MWCO). Wash the column once with PBS. Resuspend the collected protein in PBS and measure its absorbance at 280nm. (1X PBS is used as a blank.)
Chapter 3: Diary of Work Done, Results & Data Interpretation
In writing this report, the author opted to use a diary-like format: that is, a consecutive flow of the physical work done, the results of this work, and discussion of the results. There are two reasons for this:
1. A clear statement of the work done each day is the best evidence of having kept a proper diary during the project (bar handing up the rough copy, which is in no condition to correct). This is a necessary skill for any scientist.
2. By having each set of results immediately followed by the discussion, there is no need for the reader to flip back and forth between separate sections of the report.
A meeting was scheduled with Dr. Brendan O’Connor prior to the commencement of the project. This enabled the author to draw up a schematic of the work ahead, which was as follows:
Figure 6: Flow diagram of the physical work involved.
The author was also encouraged to have an attempt at primer design. This was successfully carried out by drawing upon the knowledge gained in 3rd year microbiology practicals, including how:
- The primers need to be approximately 30 base-pairs in length, with a high G-C content (more than 65%) at the 3’ end;
- The sequence of the reverse primer should be complementary to the 30 bp immediately preceding the forward primer;
- The primers should be designed such that the complementary region has a Tm of around 60˚C;
- PCR primers usually come without a phosphate group on their 5′ termini. As a result, we cannot simply ligate the ends of a PCR product together; rather, they must be phosphorylated first. This can be circumvented by ordering the primers with a phosphate already added to the 5′ end.
Preparation of agar plates –– Cultures of pQE-30 (carrying DsRed) in LBA –– Cultures of KRX and JM109 in LB
Our first task was to make up 1500 mL of LB broth (Section 2.1). To prepare for the first round of transformation on Day 7, 1200 mL of this broth was used to make up LBA agar plates (Section 2.3); the remaining 300 mL was divided into two 150 mL stocks. Ampicillin was added to one of these stocks (Section 2.2) for use in culturing our pQE-30 plasmid carrying DsRed (Section 2.5); the remaining amp-free LB was used to culture KRX and JM109 – our competent cells – to test their confluency (Section 2.4).
The vector pQE-30 (PROMEGA) flourishes in LBA, as it carries an AmpR marker. It expresses DsRed such that the protein is bound to a poly-Histidine tag, thereby enabling its purification by immobilised nickel affinity chromatography.
KRX and JM109 (PROMEGA) are high expression E. coli strains, designed to optimise transformation and the induction of protein expression. (More details are provided on Day 6.)
pQE-30 confirmation gel –– Preparation of TE Buffer –– Stocks of pQE-30 in LBA –– Stocks of KRX and JM109 in LB
Our overnight cultures of pQE-30 in LBA were a success. The plasmid was then purified from the medium (Section 2.6) and run through agarose gel electrophoresis (Section 2.7), to confirm its identity on the basis of size (3641bp). The success of this gel would enable us to proceed with mutagenesis PCR on Day 3: the induction of V105A. (The bands representing pQE-30 in lanes 2 and 3 were rather faint, indicating a low cell number; this can be put down to inadequacy in our DNA purification technique.)
TE Buffer was also made up (Section 2.8) to prepare for PCR on Day 3. Resuspending the primers in TE Buffer solubilises them, while protecting them from degradation. (Tris inactivates DNA nucleases by adjusting the pH to 8.0; EDTA does so by binding to metal cations that are required by the nucleases.)
Stocks of the pQE-30 cultures in LBA were prepared as well (Section 2.9). This would enable us to grow up the plasmid, induce the expression of DsRed, purify the protein, and compare it with our final mutant at the end of the project.
Our overnight cultures of KRX and JM109 in LB were also successful, thus demonstrating their confluency. Stocks of both strains were prepared as a precautionary measure (Section 2.9), i.e. in case problems were to arise with the competent cells to be prepared on Day 5.
Resuspension of primers in TE Buffer –– Site-directed mutagenesis, round 1/3
Before proceeding with PCR, it was necessary to resuspend the primers in TE Buffer and to make 1 in 10 dilutions of these stocks (Section 2.10). The first round of mutagenesis PCR – changing DsRed to E5up – was then carried out, with four aliquots of 50 μL for the four potential annealing temperatures: 55, 60, 65, and 70˚C (Section 2.11). On Day 4, an agarose gel would be run to confirm the success of our PCR.
PCR confirmation gel –– Repeat of PCR –– PCR confirmation gel –– DNA purification from PCR
The PCR samples were run on a 0.7% agarose gel to confirm the presence of our newly synthesised plasmid, presumably now carrying E5up. The results of this gel were poor: there was nothing in any of the lanes representing our plasmid. It is highly likely that 1 μL of template DNA was added to the PCR master mix instead of 4 μL. Thus, there was a need to repeat the PCR. The results of the second confirmation gel were as follows:
Figure 7: Results of PCR confirmation gel for pQE-30, now carrying E5up.
The plasmid is linearised; therefore, it should be more than 4000 bp in size. This is represented in lane 4 (55˚C annealing temperature). The plasmid in lane 5 is not the correct size – the result of primers binding to a different sequence. When lane 6 was loaded, the bottom of the well was pierced, causing the plasmid to leak out. The bands in lane 7 are smeared, suggesting DNA degradation. As for lanes 1 and 2, these represent unsuccessful attempts at loading the marker ladder.
Upon confirmation, the plasmid was purified from the PCR mix (Section 2.12) to prepare for a restriction digest on Day 5. The purpose of the restriction digest is to remove the template DNA and leave us with the new mutant sequence.
Restriction digest –– Preparation of TB and SOB –– Cultures of KRX and JM109 in LB –– Preparation of SOC
A restriction digest was carried out to cut away the template DNA from our purified plasmid, now carrying E5up (Section 2.13).
Terrific Broth (TB) and Super Optimal Broth (SOB) were also prepared for use in growing up our competent cells on Day 6 (Sections 2.14 and 2.15, respectively). To this end, overnight cultures of KRX and JM109 in LB were prepared as well.
The remaining SOB was used to make up SOC (SOB with catabolite repression, Section 2.16), to be used in the transformation of competent cells on Day 7.
SyBr Safe gel –– DNA purification from gel –– Ligation –– Growing up KRX and JM109 competent cells
The nicked plasmids were run on a SyBr Safe gel (Section 2.17) to confirm the success of our restriction digest and to excise the best expressed plasmid. We expected to see multiple bands representing the removed template DNA, and one ~4000bp band for the plasmid. However, this was not the case: rather, there was nothing on the first SyBr Safe gel. Given the success of the PCR, we had to conclude that mistakes were made in the restriction digest (e.g. the buffer not being mixed properly), causing the plasmid to leak out of the wells.
A faint band for our plasmid was achieved on the second attempt. We excised this band from the gel, purified the cut plasmid (Section 2.18), and sealed it up by carrying out a ligation (Section 2.19). The plasmid was now ready to be taken up and expressed by our competent cells. (If an improperly ligated plasmid is taken up and expressed by bacterial cells, the sequence is more prone to random mutation.)
The second requirement for transformation was to grow up the KRX and JM109 competent cells, using our LB cultures (Section 2.20). During the incubation step, it is important to monitor the absorbance levels of the cells, as an indicator of concentration. The optimum absorbance is 0.4 – 0.6 (read at 600nm).
The JM109 strain is more suited to transformation than KRX. Since JM109 lacks the E. coli K restriction system, recombination of cloned DNA with host chromosomal DNA is prevented. (The endonuclease A-mutation also results in an improved quality and yield of isolated plasmid DNA.) To illustrate this point, both the JM109 and the KRX aliquots would be used in the first round of transformation on Day 7.
However, since KRX is more suitable for induced protein expression (owing to the presence of a chromosomal copy of T7 RNA polymerase under the control of rhaBAD, a ramnose promoter, allowing for dramatic control over the proteins that can be expressed prior to induction), after round 3 of PCR, only the KRX transformants would be used for the induced expression and purification of our mutant protein.
Transformation of KRX and JM109 with pQE-30 carrying E5up
Our KRX and JM109 competent cells were transformed with the ligated plasmid (Section 2.21), thus enabling (and optimising) the expression of E5up – our enhanced DsRed protein. We would check for single, fluorescent colonies on Day 8. These colonies would be used to make overnight cultures in LBA, in preparation for the second round of PCR: the induction of S197T to change E5up to E5.
When we heat-shock KRX and JM109 to take up the plasmid, they gain AmpR. This is what enables them to survive on the LBA agar plates.
Cultures of JM109 expressing E5up in LBA
Single colonies were present for the JM109 plates, but not the KRX plates. While we certainly expected a higher rate of transformation for JM109 than KRX, the complete absence of KRX transformants was surprising. Given that there were only three colonies present on the JM109 plate, this can be attributed – at least in part – to our inexperience at transformation.
Figure 8: Results for transformation of concentrated JM109 with pQE-30 carrying E5up (enhanced red), viewed under the transilluminator.
Figure 8 above displays the three highly fluorescent single colonies obtained on our concentrated JM109 plate. Numerous less fluorescent colonies are also visible. In the case of the latter, it is possible that E5up is being expressed by the cells, but the protein has yet to lose most of its residual green fluorescence, thus compromising the enhanced red. It is also possible that these cells have grown on patches of agar with no amp, and are now dying – the result of not mixing the amp sufficiently with the agar.
Figure 9 below sheds more light on the situation, and also serves as a warner against loading two different fluorescent proteins on the same SyBr Safe gel:
Figure 9: Contamination of GFP with DsRed (expressed by JM109 and/or KRX).
Colonies of JM109 and/or KRX expressing E5up are visible among the mass of yellow-green colonies (presumably expressing EYFP) belonging to colleague Gary Gillick. Since we shared the same SyBr Safe gel, it is safe to conclude that leakage from our wells settled in his. Undoubtedly, this loss of plasmid concentration played a role in the scarcity of fluorescent single colonies on our plates. To guard against cross-contamination in the future, we would make sure to run our digests on separate gels.
Proceeding with the assumption that we had obtained single colonies of JM109 expressing E5up, overnight cultures were prepared in LBA. The plasmid would be purified on Day 9, to prepare for round two of PCR.
DNA purification from LBA –– Site-directed mutagenesis, round 2/3 –– Stocks of JM109 expressing E5up in LBA
Under the transilluminator, our overnight cultures glowed bright red; thus, we had achieved 5 mL cultures of JM109 expressing E5up. The plasmid was subsequently purified and used for the second round of PCR: the induction of S197T to change E5up to E5.
As was done for DsRed, stocks of JM109 expressing E5up were prepared for comparing our mutants at the end of the project.
PCR confirmation gel
The plasmids were run a 0.7% agarose gel, to confirm the success of our PCR. The results were as follows:
Figure 10: Results of PCR confirmation gel for pQE-30, presumed to be carrying E5.
A band of approximately 4000bp can be seen in lane 5 (70˚C annealing temperature). This was the only band on the gel; the smear in lane 2 (representing millions of copies of plasmid DNA) tells us that the 55˚C sample was degraded.
We wanted the maximum amount of E5 to work with, so we needed to be reasonably certain that only the 70˚C sample contained the newly synthesised plasmid. To this end, a 40 minute gel was run with double the number of samples. The results of this gel were as follows:
Figure 11: Results of 40-minute PCR confirmation gel for pQE-30, with twice the number of samples.
Once again, the only bands that appeared were for the 70˚C sample (lanes 5 and 10). Thus, we would proceed with DNA purification from PCR tube 4 on Day 11.
Days 11 & 12
DNA purification from PCR mixture –– Restriction digest –– SyBr Safe gel –– DNA purification from gel –– Ligation –– Preparation of agarose gels
These two days were spent preparing for the second round of transformation. Upon purification of pQE-30 – now carrying E5, presumably – from the PCR mix, a restriction digest was carried out. The resulting digests were run on a SyBr Safe gel, with the best expressed plasmid being excised, purified and ligated. The plasmid was now ready to be taken up and expressed by our competent cells on Day 13.
There was a worry that the elution buffer in the miniprep kit was at fault. Thus, it was decided that distilled water would be used from now on.
Three agarose gels were also made up for future use. These gels were stored at 4˚C with a small amount of 1X TAE buffer, to keep them from drying out.
Transformation of JM109 with pQE-30 carrying E5 –– Preparation of SOC
Our JM109 competent cells were transformed with the ligated plasmid, to achieve the optimal expression of E5. For the reasons given on Day 6, only the JM109 aliquots were used this time; the KRX aliquots would be used for the third round of transformation and subsequent protein expression analysis.
For the concentrated plates, given the large bands obtained for our plasmid on Day 10 (second PCR gel), it was decided to prepare a 1 in 4 dilution, to ensure the presence of single, fluorescent colonies on Day 14.
It was necessary to make up SOC (from 100 mL of SOB) before proceeding with the transformation, as the previous stock had become contaminated. SOC contains more nutrients than LB, and thus has a shorter shelf life.
Just before proceeding with the spread plates, however, it was discovered that all of our LBA agar plates had been contaminated with fungus. Given that the plates were prepared under sterile conditions and kept at 4˚C, it is likely that the agar simply wasn’t allowed to cool down sufficiently when amp was first added, thus negating its effects and allowing for eventual contamination from a variety of sources.
Donal Moynihan, a PhD student at the ISSC, permitted us to use some of his plates instead. We would return the favour on Day 14 by making up fresh plates.
Preparation of SOC –– Preparation of agar plates
Not one colony was present on anyone’s plates. While it is common knowledge that rates of success for transformation decrease after each round, once again, the complete lack of transformants was surprising. Probability dictates that something all three of us were working with was at fault, rather than all three of us making serious errors in the transformation. This may well have been the SOC broth, as it was not mixed properly in its preparation (the heat-stir wasn’t used). This was made up again to repeat the transformation on Day 15.
20 fresh plates of LBA agar were also prepared, to replace the ones given to us on Day 13.
Transformation of JM109 with pQE-30 carrying E5 –– Streak plates of JM109 expressing E5up
We repeated the transformation of our JM109 competent cells with pQE-30, presumed to be carrying E5.
In addition, the one fluorescent colony of JM109 expressing E5up obtained on our neat plate (Day 8) was used to streak for single colonies (Section 2.22). On Day 16, these colonies would be used to make overnight cultures in LBA, from which we would purify our plasmid to send for sequencing. We needed to be absolutely certain that the first mutation was induced; for if it was not, then proceeding to induce the other mutations required to obtain AG4 would be pointless.
Transformation of JM109 with pQE-30 carrying E5up –– Culturing pQE-30 carrying E5up in LBA
Once again, there were no colonies on the plates for any of us. To narrow down the possibilities of what went wrong, it was decided to repeat the first transformation, i.e. the uptake and expression of pQE-30 (carrying E5up) by KRX and JM109. We reasoned that if this worked – and there was no reason to think that it wouldn’t, as it worked the first time – then there were three potential issues:
1. Something went wrong in preparing E5 for transformation – possibly the ligase becoming faulty;
2. The plates were at fault (unlikely, as they were prepared by the book);
3. The SOC was at fault (unlikely, as extra care was taken in its preparation).
There was a need to make up more LBA agar plates before proceeding with the transformation. It was also decided that LB broth would be used instead of SOC from now on, as it is less prone to contamination. We would check for single colonies on Day 17.
On our E5up streak plate, there was an abundance of fluorescent single colonies. These were used to inoculate overnight cultures of LBA. This would be followed up with a plasmid prep on Day 17.
Ligation –– DNA purification from LBA
Yet again, nothing appeared on our plates. This was surprising, as our previous E5up transformation worked just fine; clearly there was nothing wrong with our ligated plasmid. Neither should there have been anything wrong with our competent cells, as they had been kept at -80˚C the whole time. The ligase becoming faulty may explain the failure of our E5 transformation attempt, but it doesn’t explain this; clearly there was nothing wrong with the ligase originally. There was no issue with our plates either (no contamination, agar prepared in separate 500 mL jars for each of us) or our LB, for the same reasons.
At the time, we simply had to be content with this gap in our understanding. All we could do was proceed with the assumption that our plasmid (carrying E5) wasn’t ligated properly. There was just enough of the purified E5 digest left over; thus, we performed a ligation (with a fresh tube of ligase, to rule out the possibility of it being an issue next time) and would attempt transformation again on Day 18.
We also purified pQE-30 (carrying E5up) from the cultures in LBA. The purified ligation would be used for PCR on Day 18, to provide us with E5 in case the new transformation doesn’t work. We would also send the ligation for sequencing on Day 19.
Transformation of JM109 with pQE-30 carrying E5 –– Site-directed mutagenesis, round 2/3 (repeat)
Using our fresh E5 ligation, the transformation of JM109 was attempted once more.
By this time, we had concluded that since the failure of our second E5up transformation had nothing to do with the ligase, clearly there was some issue with our competent cells. Perhaps the aliquots used for the second E5up transformation were faulty, whereas those used for the first E5up transformation were just fine.
We reasoned that with our newly ligated plasmid (carrying E5), if the transformation still doesn’t work, then it would be safe to conclude that our competent cells have been compromised. In this case, we would use Donal’s competent cells for transformation on Day 20, using the plasmid to be obtained from PCR on Day 19 (carrying E5).
On the other hand, if the transformation does work, then at least some of our competent cell stocks are fine. In this case, we would make overnight cultures on Day 19 and proceed with the third round of PCR (inducing V71M to obtain AG4) on Day 20.
The second round of PCR was repeated for some of our first ligation (carrying E5up), to provide us with a backup of E5. The remainder would be sent for sequencing on Day 19.
PCR confirmation gel –– DNA purification from PCR mixture –– Restriction digest –– SyBr Safe gel –– DNA purification from gel –– Ligation –– Sending plasmid DNA for sequencing
The transformation didn’t work, thus demonstrating that our JM109 cells were no longer competent. Having been provided with fresh competent cells by Donal, we prepared our newly synthesised plasmid (carrying E5) for what would hopefully be the final attempt at transformation on Day 20.
The plasmids were first run on an agarose gel, to confirm the success of our PCR. The results were as follows:
Figure 12: Results of PCR confirmation gel for pQE-30, presumably now carrying E5.
Bands of around 4000bp could be seen for the 65 and 70˚C samples, thus indicating the presence of E5. The double band obtained in lane 4 suggests the presence of a large contaminant with our plasmid; thus, we only combined the 65 and 70˚C samples.
After this, we purified the plasmid from the PCR mix, performed a restriction digest, and ran the digests on a SyBr Safe gel. The most concentrated band was excised, and the DNA was purified and sealed up with a ligation.
Our purified ligation from Day 18 (carrying E5up) was sent for sequencing (Section 2.23).
Transformation of JM109 with pQE-30 carrying E5
The new aliquots of JM109 were transformed with our freshly ligated plasmid, carrying E5. We reasoned that if this worked, then we would know with absolute certainty that our competent cells were the problem.
Streak plates of JM109 expressing E5
At long last, fluorescent single colonies appeared for our E5 transformation. We could thus say with certainty that the competent cells were at fault. Perhaps they were allowed to thaw out for too long in our first attempt.
Figure 13: Results for transformation of JM109 with pQE-30 carrying E5 (enhanced red), viewed under the transilluminator.
Five of these colonies were used to prepare streak plates, to supply us with an abundance of colonies for making overnight cultures on Day 23.
Cultures of JM109 expressing E5 in LBA
Five LBA samples were inoculated with single colonies of JM109 expressing E5. On Day 23, we would purify our plasmid from the best expressed clone, and proceed with round three of PCR: the induction of V71M to obtain green-emitting AG4.
Stocks of JM109 expressing E5 –– DNA purification from LBA –– Site-directed mutagenesis, round 3/3 –– PCR confirmation gel –– Sequencing results for E5up
Our LBA samples were successfully inoculated. As was done for DsRed and E5up, stocks of JM109 expressing E5 were prepared for comparing our mutants at the end of the project. The plasmid was then purified from the LBA, and used for the third round of PCR. Once the PCR had run its course, the newly synthesised plasmid (presumably now carrying AG4) was run on a 0.7% agarose gel.
According to the results (no image available), annealing and extension occurred for our 60, 65 and 70˚C samples. As before, however, there was a concern that the band for our 65˚C sample might consist of two plasmids; therefore, only the 60 and 70˚C samples were combined. We would proceed with DNA purification on Day 24.
As a side note, here is what happens when DNA samples are run on an agarose gel that hasn’t set properly:
Figure 14: The results of agarose gel electrophoresis when the agarose hasn’t set properly.
Although the bands are all of the same size, the incomplete solidification of the gel has affected their positions. (This happened to us at least once.) It is important to take such basic considerations into account when faced with data that doesn’t appear to make sense.
The sequencing results had finally arrived for E5up. The data gave us the same exact sequence as DsRed, but with one crucial difference: at amino acid 105, there is an Alanine instead a Valine. This is highlighted in blue below:
Figure 15: The amino acid sequence of DsRed-E5up (EXPASY translation, forward primer) showing the replacement of Valine with Alanine at position 105.
Our primers were designed to incorporate a Cysteine instead of a Thymidine at nucleotide 314, to alter GTG to GCG; this results in Alanine instead of Valine. Thus, we had successfully induced the first of three mutations required to obtain AG4.
DNA purification from PCR mixture –– Restriction digest –– SyBr Safe gel –– DNA purification –– Ligation
Our newly synthesised plasmid – presumed to be carrying AG4 – was purified from the PCR mix. This was followed by a restriction digest, and the digests run on a SyBr Safe gel. The most concentrated band was excised, and the nicked plasmid was purified and ligated. This left us in a prime position for transformation on Day 25.
Transformation of JM109 with pQE-30 carrying AG4
The JM109 cells provided to us by Donal were transformed with our newly ligated plasmid. We reasoned that if green fluorescent colonies appeared on our plates, then we could be reasonably sure that V71M was successfully induced.
Streak plates for red and non-fluorescent colonies of JM109 –– Transformation of JM109 with pQE-30 carrying AG4 –– Ligation
There were a few red fluorescent colonies on the concentrated plate, as well as an abundance of colonies displaying no fluorescence (no image available). We reasoned that either the mutation worked, or it did not. If the mutation did not work:
1. Red colonies are E5 (or E5up), non-fluorescent colonies are faulty E5 (or E5up);
2. Red colonies are contamination from E57, non-fluorescent colonies are faulty E5 (or E5up).
The induction of S197T does not alter the red fluorescence, so there is no visible difference between E5up and E5. Conversely, if the mutation worked:
1. Non-fluorescent colonies are AG4, red colonies are contamination from E57;
2. Non-fluorescent colonies are AG4, red colonies are contamination from E5 (or E5up).
Knowing that the induction of V71M negates the enhanced red fluorescence, and assuming that we had induced S197T, we speculated that perhaps the green fluorescence is only visible under UV light; we would check this after purifying the protein by IMAC. Sequencing would confirm the protein’s identity long before this.
Streak plates were prepared for both the red and non-fluorescent colonies, thus enabling the preparation of overnight cultures on Day 27.
For the purposes of protein expression analysis, the AG4 transformation was also repeated using KRX. LBA and TBA were made up to this end.
Our purified ligation (carrying E5) was also sent for sequencing.
Cultures of red and non-fluorescent colonies of JM109 in LBA and TBA –– Preparation of SDS Polyacrylamide gel –– Streak plates of KRX colonies
Cultures of both the red and non-fluorescent colonies obtained for our Day 26 streak plates were prepared in LBA and TBA (Section 2.24). On Day 28, the TBA cultures would be used to identify our best expressed clone via SDS-PAGE. The LBA cultures would then be used for the induced expression and purification of this clone. As explained on Day 6, the latter would only be done for KRX; this would make the JM109 gel a practice run, so to speak.
An SDS Polyacrylamide gel was also prepared for this purpose (Section 2.25), and stored overnight at 4˚C. (Heating the samples in SDS buffer coats the proteins in a negative charge. As a result, when the gel is run, all the proteins move towards the positively-charged electrode, meaning they separate on the basis of size, rather than charge. This enables the identification of any potential contaminants.)
Streak plates were also prepared using the colonies obtained for our KRX transformation. The resulting single colonies would be used to make overnight cultures on Day 29.
Stocks of M3 Red and M3 Green in LBA –– SDS-PAGE for M3 Red and M3 Green
The cultures of non-fluorescent colonies prepared in LBA and TBA were now fluorescing yellowish-green; we presumed this was AG4. The cultures of red colonies were now super-red; we presumed this to be E5 (or E5up), or possibly contamination from colleague Andrius’ E57 plasmid (also enhanced red). From now on, these proteins would be referred to as M3 Green and M3 Red respectively.
Our cultures for M3 Red and M3 Green (JM109) in LBA appeared as follows:
Clearly we had a different fluorescent protein for M3 Green, though perhaps not AG4; the sequencing results would reveal all. Stocks of both were prepared for this purpose.
The difference between our overnight cultures in TBA was less striking. However, by centrifuging 1 mL of each to pellet the cells, it was possible to confirm their different fluorescence patterns.
Therefore, we proceeded to use both of these cultures for SDS-PAGE (Section 2.26), with the gel to be de-stained on Day 29. Ideally, this would enable the identification of our best expressed clone, as would be done for KRX on Day 31.
However, the gel didn’t run properly, as the gaskets were left on: rubber insulates against electricity. Thus, the gaskets were removed, more SDS-PAGE buffer was added, and the gel was rerun. It would become clear how this affected the protein on Day 29.
DNA Purification from glycerol stocks (error) –– Sending M3 Red and M3 Green for sequencing –– Streak plates for KRX single colonies
Unsurprisingly, the de-stained JM109 gel appeared distorted:
Figure 17: Distorted SDS Polyacrylamide gel. Proteins do not separate out well unless they are forced to migrate by an opposing electrical charge. The rubber gasket, if left on, acts as a barrier between the proteins and this charge.
Bands were present for our proteins, but it was impossible to tell whether or not they were the correct size (~28 kDa). Thus, we would exercise particular caution in readying our KRX gel on Day 31.
Using the stocks made on Day 28, our ligations carrying M3 Red and M3 Green were purified and sent for sequencing. This was a mistake, as the cells need to cultured in LBA first: up to a third of the cells stored at –80˚C in glycerol die straight away. Thus, we expected the sequencing results to be inconclusive.
As regards the KRX streak plates prepared on Day 27, although single colonies were present, there was not an abundance of them; this was due to poor streaking technique. Thus, five more plates were streaked in the correct manner, with the resulting colonies to be used for the preparation of overnight cultures on Day 30.
Cultures of KRX expressing M3 Red and M3 Green in LBA and TBA –– Preparation of SDS Polyacrylamide gel
Using the colonies obtained on our new KRX streak plates, overnight cultures of KRX expressing M3 Red and M3 Green were prepared in LBA and TBA, for protein screening on Day 31.
An SDS Polyacrylamide gel was also prepared for this purpose, to be stored overnight at 4˚C.
SDS-PAGE for M3 Red and M3 Green –– Cultures of KRX expressing M3 Red and M3 Green in LBA and TBA
The TBA was successfully inoculated; thus, we proceeded with SDS-PAGE for KRX expressing M3 Red and M3 Green.
The cultures in LBA, on the other hand, were contaminated. Thus, fresh LBA was made up for a redo of our overnight cultures. Following the identification of our best expressed clone, these cultures would be used to inoculate 200 mL of TBA, for the induction of protein expression.
Cultures of KRX expressing M3 Red and M3 Green in LBA and TBA –– Preparation of SDS Polyacrylamide gel
On the de-stained KRX gel, the correct-sized bands were visible for M3 Red (lanes 3-5), with our best expressed clone in lane 3. However, there were no definitive bands for M3 Green (lanes 6-7). The bands that were present in lanes 6-7 were not the correct-size (~28 kDa), thus, we could not proceed to induce the expression of M3 Green. (No image available.) We reasoned that it was possibly a partially-ligated plasmid, and thus was not expressing as it should have been; or perhaps poor loading technique was the source of the problem.
At this point, the sequencing results had yet to arrive for either plasmid; thus, on Day 33, SDS-PAGE would be repeated for both M3 Green and M3 Red, for the sake of consistency. This necessitated making up new overnight cultures in TBA and LBA, as well as a new SDS Polyacrylamide gel.
SDS-PAGE for M3 Red and M3 Green –– Stocks of KRX expressing M3 Red and M3 Green in LBA
Using the overnight cultures prepared in TBA, KRX was rerun through SDS-PAGE for M3 Red and M3 Green. The gel would be de-stained on Day 34.
Assuming that the correct-sized bands (~28 kDa) would be present for both proteins, stocks were made from our cultures of KRX in LBA.
Cultures of KRX expressing M3 Green and M3 Red in LBA
The de-stained gel for KRX was identical to the previous one: sizeable bands appeared for M3 Red (some bulkier than others), while no definitive bands appeared for M3 Green.
Just in case the bands for M3 Green were not telling the whole story, on Day 35, we would induce the expression of both proteins. Before doing so, however, there was a need to make up new cultures in LBA, using the stocks prepared on Day 33.
A note on induced protein expression
Along with obtaining a high concentration of protein, one reason that we always try to keep the environment induced is the susceptibility of Methionine residues to oxidation – a problem when we have very pure solutions of protein. As the Methionine becomes modified, the result is a misfolded protein response. This oxidation occurs often, and is a key problem in large-scale protein purification. In fact, there is a large focus on engineering out this Methionine in protein drugs, as modified Methionines can sometimes lead to an antigenic protein.
Preparation of TBA –– Stripping and recharging the IMAC resin –– Inducing the expression of M3 Red and M3 Green –– Preparation of lysis buffer
Using the KRX cultures prepared in LBA on Day 34, we proceeded to induce the expression of M3 Red and M3 Green, using IPTG (Section 2.27).
There was a need to make up 400 mL of TBA before proceeding. There was also a need to strip the IMAC resin of any nickel ions to which proteins may be bound, and recharge it with 100mM NiSO4 (Section 2.29).
IPTG is an acronym for Isopropyl β-D-1-thiogalactopyranoside, a synthetic analogue of lactose that is highly stable. IPTG inactivates the lac repressor in E. coli and promotes lactose utilisation by inducing the synthesis of beta-galactosidase. IPTG is thus used to induce the expression of cloned genes that are under control of the lac operon in E. coli.
After the addition of IPTG, it takes approximately two hours for the KRX cells to reach an absorbance of 0.4 – 0.6. After 2.5 hours, our KRX cells expressing M3 Red had an absorbance of 0.562. However, the cells expressing M3 Green only had an absorbance of 0.093, which absolutely does not indicate cell growth. We reasoned that our KRX cells expressing M3 Green had become contaminated: after all, the aliquots used for absorbance measurement were not taken in the fume-hood, as they should have been. This being the case, we would postpone obtaining M3 Green protein until Day 44, and continued with the expression and purification of M3 Red.
On Day 36, we would obtain M3 Red protein using a cell disruptor, and then purify it via IMAC. To prepare for the latter, various concentrations of Lysis buffer were made up (20, 80 and 250mM imidazole). (See Section 2.28.)
Lysis of KRX cells to release M3 Red protein –– Protein purification by IMAC –– Absorbance
On Day 35, once our induced KRX cells had reached the optimum absorbance, the LBA was left to incubate at 30˚C for 16 hours. On this day, the cells were lysed to obtain M3 Red (Section 2.30), which we proceeded to purify by IMAC (Section 2.31). The filtered lysate, the unbound lysate, the various washes, and the protein elutions were all collected. On Day 37, we would be able to assess the purity of our fractions via SDS-PAGE.
Our most concentrated M3 Red elution appeared as follows under the transilluminator:
Figure 18: M3 Red protein (presumed to be E5up, E5, or E57) in lysis buffer (500 mM imidazole) after purification by immobilised nickel affinity chromatography. Evidently, the maturation of the DsRed fluorophore has reached its conclusion by the induction of V105A.
However, the brightness of our elutions was no guarantee that they were free of any major contaminants. This would become clear upon reading the absorbance on Day 38.
A note on IMAC
Our affinity column contains a Nitrilotriacetic acid resin, charged with Ni2+. Once the filtered lysate for M3 Red is added, the protein’s high affinity for Ni-NTA allows for the poly-Histidine tag to bind to the immobilised nickel ions on our resin. Most of the contaminant proteins in the added sample are washed through the column; those that bind have a weaker affinity for the resin, and are thus washed away using 20, 50, 80 and 100mM concentrations of imidazole buffer. M3 Red is then eluted by simply using a higher concentration of imidazole (200-250mM), as this causes the displacement of the His-tag.
As for the tag itself, this consists of six Histidine residues in tandem, with an enterokinase cleavage site to allow its ultimate removal. Six seems to give us the optimal affinity binding to immobilised nickel. Histidines carry negative charges at neutral pH levels (7.0-7.4); thus, with six Histidines in tandem, we have a poly-anionic tag that has a very high affinity for a divalent cation like nickel – higher even than that of antibody for antigen, or enzyme for substrate. Binding of the His-tag does not depend on the 3-D structure of the protein; rather, even if the tag is inaccessible, it will bind so long as two or more Histidines are available to interact with the nickel cation.
SDS-PAGE for M3 Red
Our M3 Red fractions were run through SDS-PAGE, to investigate the presence of impurities. Any residual cell debris should have eluted in the unbound lysate; any major contaminant proteins should have eluted in the 20 mM and 80 mM imidazole washes. The gel would be de-stained on Day 38, thus giving us our answer.
Buffer exchanging M3 Red in PBS –– Reading absorbance –– Fluorescence under UV light
The de-stained gel provided us with the ideal results:
1. Most of the contaminant proteins were eluted in the imidazole washes, though some remained in the M3 Red elutions.
2. The 28 kDa band obtained for our protein elutions (lanes 3-9) was present for the filtered lysate (lane 10, circled in red), but was absent for the unbound lysate (lane 11) and imidazole washes (lanes 12 and 13). This indicates that virtually all of the protein ended up binding to the column, remained bound during the washing steps, and was only eluted with the addition of 500 mM imidazole lysis buffer.
Figure 19: The results of SDS-PAGE for our M3 Red elutions, the filtered lysate, the unbound lysate, and the imidazole washes.
The enduring presence of contaminants necessitated buffer exchanging our fractions into 1X PBS (Section 2.32); this was done for our most concentrated samples. Any contaminant proteins were expected to be resolved here. However, according to the absorbance readings, this was not to be; rather, a second large peak appeared on the spectrophotometer:
Figure 20: Absorbance readings for our buffer-exchanged M3 Red elutions. The presence of another tall peak at ~650nm suggests that a large unidentified protein has remained in the elution.
This indicated the presence of another large protein in our samples, a protein that somehow eluded the washing steps. (This was not an imidazole peak, as PBS would have washed it out.) We posited that this was a subunit of our protein, but it was difficult to know without any sequencing results at hand. Without them, there was no option but to repeat the induced expression of M3 Red, and purify it again via IMAC. A gradient imidazole wash would be used this time, to completely remove any and all contaminant proteins.
On Day 39, we would prepare overnight cultures of M3 Red and DsRed in LBA, using the stocks in our possession. Prior to this, we were to run our buffer-exchanged fractions through SDS-PAGE to confirm the presence of this suspected contaminant protein and, if possible, identify it. (The purpose of growing up and purifying DsRed is to be able to demonstrate a shift in absorbance from our original protein to our mutant, representing the enhanced fluorescence.)
Our samples were also viewed under UV light. Our M3 Red elution exhibited a considerably more pink fluorescence than what was presumed to be E57 (enhanced red):
Figure 21: Our M3 Red elution (left) compared to presumed E57 (right) under UV light. The sequencing results would confirm if this amounted to an actual difference in nucleotides.
SDS-PAGE of buffer-exchanged M3 Red –– Cultures of KRX expressing M3 Red & JM109 expressing DsRed in LBA –– Sequencing results for M3 Red and M3 Green –– Cultures of KRX expressing M3 Green in LBA
Our buffer exchanged M3 Red fractions were run through SDS-PAGE. De-staining the gel on Day 40 would allow us to identify the contaminant protein responsible for the second absorbance peak. Given the concentration of our sample, dilutions to 1 in 6 were prepared and run on the gel alongside the neat protein.
As the gel was running, however, the power station kept switching between volts and amps. This was only noticed approximately 45 minutes after the beginning of electrophoresis, so this may have been the case from the start. There was a concern that this stop-start behaviour could cause the fractions to be smeared or appear at different positions on the gel. This would make it difficult to identify the contaminant protein; we would find out on Day 40.
Assuming the presence of this contaminant protein, on Day 40, we would repeat the induced expression of M3 Red, and also induce the expression of DsRed. To prepare for this, overnight cultures of KRX (expressing M3 Red) and JM109 (expressing DsRed) were prepared in LBA.
The sequencing results had finally arrived for M3 Red and M3 Green (nothing yet for E5). However, as expected, the results were inconclusive: we only received half of the sequence for M3 Red, and nothing for M3 Green (“ERROR”). Undoubtedly, this is because the plasmids were not grown up in LBA before being purified and sent off: rather, they were merely purified from the glycerol stocks. The plasmids need to be given a chance to be expressed, as explained on Day 29.
In accordance with this, an overnight culture of KRX expressing M3 Green was prepared in LBA. On Day 40, we would perform a plasmid prep for both M3 Green and M3 Red, and send them for sequencing once again.
Inducing the expression of M3 Red & DsRed –– DNA purification from LBA –– Sending M3 Red and M3 Green for sequencing –– Confirmation gel
De-staining the gel from Day 39 confirmed our fears: the protein fractions ended up at different positions on the gel, and the ladder was smeared. Therefore, it was futile to try and identify the contaminant protein. Repeating the process of induced expression and IMAC to obtain a purer protein sample was considered a more worthy pursuit. On Day 42, when running our new protein elutions through SDS-PAGE, we would make sure to monitor the power station very closely.
On Day 39, our LBA samples were successfully inoculated with KRX (expressing M3 Red) and JM109 (expressing DsRed). Thus, having made up fresh TBA, we proceeded to induce the expression of both proteins. On Day 41, we would lyse our KRX cells to release the proteins, and purify them by IMAC.
Our cultures of KRX for M3 Green and M3 Red were also purified, and sent for sequencing. Given our previous sequencing results, however, it was prudent to run the purified plasmids on an agarose gel first. The results of this gel were as follows:
Figure 22: Results of a 0.7% agarose gel to confirm the presence of pQE-30 carrying M3 Red and pQE-30 carrying M3 Green.
While the reader will note that the gel split in two, he will also note the presence of correct-sized bands (<4000bp) for the plasmid in all six lanes (2-7). (The bands in lanes 2 and 3 represent a small amount of plasmid that didn’t leak out. 50% Glycerol was added to allow the samples to settle in lanes 4 to 7.) Thus, we could rest assured that KRX was expressing the proteins adequately, and thus expect working sequence data.
Lysis of KRX to release M3 Red and DsRed –– Protein purification by IMAC
Our KRX cells were lysed to release M3 Red and DsRed. A gradient imidazole wash (20, 50, 80 and 100mM) was then used to elute any and all unwanted proteins. We would check the absorbances of our fractions on Day 42.
As expected, our elutions for M3 Red were considerably brighter than those for our original DsRed:
Figure 23: M3 Red (super-enhanced DsRed, left) compared to wild-type DsRed (right) under the transilluminator.
This result, if nothing else, demonstrates the power of mutation: how just one or two single base-pair changes can dramatically affect the phenotype of a given protein.
Checking absorbance –– SDS-PAGE for M3 Red
Our red fluorescent protein was assumed to be either E5 (or E5up) or E57. On a spectrophotometer, the altered emission pattern that results from changing DsRed to E5up is represented by a diminished peak at 483nm (among the other 6 peaks):
Figure 24: Fluorescence excitation and emission spectra of matured (~2 weeks at 25°C) compared to E5up. Note the diminished peak at 483 nm in the excitation spectrum after prolonged maturation. Image sourced from Terskikh et al. (2001).
By inducing the expression of DsRed and purifying it in tandem with M3 Red, we had hoped to be able to demonstrate this shift in absorbance. However, our results for DsRed were very poor. It could be that most of the protein was eluted in the washing steps:
Figure 25: Absorbance readings for original DsRed elutions. These results would indicate that DsRed is no longer present in the solution, i.e. serious mistakes were made in the elution steps.
However, a positive result was achieved for M3 Red:
Figure 26: Absorbance readings for our second series of M3 Red elutions. The repeat appearance of this large unknown peak – particularly after a gradient imidazole wash – prompted us to think again about this protein’s identity.
As with the results obtained on Day 38, these readings compared quite favourably with those obtained by Terskikh et al., thus reinforcing our position that the protein was either E5 (or E5up) or E57. Once again, however, there was a second peak present, similar in size to that of our protein. To confirm the presence of this unknown protein, our fractions were run through SDS-PAGE. The gel would be de-stained on Day 43.
We did not have enough time to repeat the induced expression and purification of DsRed. This doesn’t matter, however, as we know that we had DsRed to begin with; this makes obtaining the correct absorbance little more than a formality.
Checking absorbance –– Buffer exchanging M3 Red in PBS –– Checking absorbance –– Cultures of KRX expressing M3 Green in LBA
Our de-stained gel confirmed the presence of this large contaminant protein:
Figure 27: Results of SDS-PAGE for neat M3 Red (lane 2) and dilutions to 1 in 6 (lanes 3-7). The protein ladder did not come out well; thus, it was impossible to positively identify the large unknown proteins in each lane as dimers of M3 Red.
Thus, we buffer exchanged our M3 Red fractions in PBS and read their absorbances again. The results remained the same, however: the second peak was unaffected. Given the size of this protein, as well as the lengths to which we had purified our samples – a gradient imidazole wash to remove all proteins that were not His-tagged, along with buffer exchanging in PBS – we reasoned that it was unlikely to be anything other than a dimer or subunit of DsRed. In fact, according to Campbell et al. (2002), DsRed and its mutants have a tendency to dimerise and still fluoresce. Surely, then, the large bands on our gel could simply be dimers of M3 Red (approximately 5600bp).
However, the ladder on our gel did not come out very well: this made it difficult to be sure. Therefore, on Day 45, the gel would be rerun with two ladders.
Overnight cultures of KRX expressing M3 Green were also prepared in LBA. On Day 44, we would use these cultures to induce the expression of M3 Green.
Inducing the expression of M3 Green –– Preparation of TBA –– Preparation of SDS Polyacrylamide gel
Using our successfully inoculated LBA samples, we proceeded to induce the expression of M3 Green. There was a need to make up 400 mL of TBA before proceeding. We also prepared an SDS Polyacrylamide gel for Day 45, in order to assess the purity of our M3 Green fractions.
Sequencing results for M3 Green and M3 Red –– Mutagenesis PCR, round 3/3 (repeat) –– Preparation of SDS Polyacrylamide gel
The sequencing results arrived for M3 Green and M3 Red, as well as for E5. The results for M3 Red indicated that we successfully induced V71M and V105A (highlighted in green), but not S197T (highlighted in red):
Figure 28: The amino acid sequence for M3 Red (EXPASY translation, reverse primer – reverse complement), demonstrating the induction of V71M and V105A, but not S197T.
1. As we explained on Day 23, the Alanine at amino acid 105 represents the induction of V105A.
2. The Methionine at amino acid 71 represents the induction of V71M. Our primers were designed to incorporate an Adenosine instead of a Guanine at nucleotide 211, to alter GTG to ATG; this results in Methionine instead of Valine.
3. The enduring presence of Serine at amino acid 197 reflects the absence of S197T. The induction of S197T changes Serine to Threonine: our primers were meant to incorporate an Adenosine instead of a Thymine at nucleotide 589, to alter TCC to ACC.
This suggested that our attempt at inducing S197T was a failure. However, the sequencing results for E5 indicated something else: that without knowing it, there never was any attempt to induce S197T. As shown below, V71M (highlighted in green) was present for our second mutant:
Figure 29: The amino acid sequence for our second mutant (EXPASY translation – forward primer), showing the induction of V71M instead of S197T. The induction of V105A is highlighted in blue.
Given that V71M was meant to be our third mutation, this suggests that we induced V71M twice, without ever trying to induce S197T. We have to put this down to human error: perhaps the primers were mislabelled. This means that our second and third mutants are the same: they both carry V105A and V71M, but not S197T. Since there is no official name for this mutant, we will simply deem it “E5up with V71M”.
For DsRed, the induction of V71M, having previously induced V105A and S197T, is meant to trigger a change from red to green fluorescence. It could be that when V71M is induced in E5up without having first induced S197T, rather than the enhanced red being negated, it is merely altered slightly. If so, this would explain what was observed on Day 38: that E5up with V71M has a more pinkish fluorescence than regular E5up. (The latter is what Andrius’ protein has turned out to be, and not E57.)
The sequencing results for M3 Green were unclear. As with M3 Red, we could see that V71M and V105A were induced (highlighted in green and blue, respectively), but not S197T (highlighted in red):
Figure 30: Part of the amino acid sequence for M3 Green (EXPASY translation – forward primer), demonstrating the induction of V71M and V105A, but not S197T.
However, there were at least four random mutations present as well. At amino acid positions 67-69, we are supposed to see Tyrosine, Glycine, and Serine (YGS) sitting just before the Lysine (K) that precedes the sixth Methionine. These have been highlighted in yellow for M3 Red below:
Figure 31: The amino acid sequence for M3 Red, with special emphasis on the residues preceding the Methionine at position 71.
In M3 Green, however, instead of YGS, we see Proline, Arginine, Cysteine and Threonine (PRCT). Thus, not only have Tyrosine, Glycine, and Serine been altered at random, but a fourth amino acid (Threonine) has been introduced, presumably via a frameshift mutation. Once again, this has been highlighted in yellow:
Figure 32: Part of the amino sequence for M3 Green, showing random mutations at positions 67-70.
It is also worth noting that the full sequence is nearly divided in half over two reading frames:
Figure 33: The complete amino acid sequence for M3 Green, divided across two reading frames.
With these abnormalities and with the absence of S197T, clearly this protein is not AG4: rather, it is likely to be a partially-ligated version of E5up with V71M, and is thus not worth purifying. Therefore, we would not proceed with cell lysis, IMAC or SDS-PAGE for M3 Green. Instead, using the time we had left, we would attempt to obtain AG4 by inducing S197T.
Mutagenesis PCR was carried out on this day, with a confirmation gel to be run on Day 46. Another SDS Polyacrylamide gel was also prepared, thereby enabling us to run two gels on Day 46 for E5up with V71M. Ideally, the first gel would show that all proteins similar in size to DsRed in the filtered lysate were indeed eluted, and are thus not represented by the large bands obtained on Day 43. (We should have run this before.)
The second gel would be used to rerun our neat protein and dilutions (to 1 in 3). This was necessary, as our ladder did not come out well on Day 43, thus preventing us from positively identifying the large protein as a dimer of DsRed.
PCR confirmation gel –– DNA purification from PCR mixture –– Restriction digest –– SyBr Safe gel –– DNA purification from gel –– Ligation –– SDS-PAGE
The correct-sized bands were present for each annealing temperature, thus confirming the success of our PCR. Presumably, the plasmid now carries AG4. We could have achieved this earlier, had not we not wasted time inducing V71M again – that is, had we received the sequencing results for our second mutant on time.
We proceeded to purify our plasmid, carry out a restriction digest, run the digests on a SyBr Safe gel, excise and purify our best expressed plasmid, and perform a ligation. The ligated plasmid was then left with Jonathan and Donal for transformation at a later date, for by this point, we had exceeded the time allocated for the project.
SDS-PAGE was also conducted for the purposes described on Day 45. We would de-stain both gels on Day 47.
On the first de-stained gel, we could see the presence of contaminant proteins in our lysate and washes, some rather large, thus bolstering the view that the unidentified bands observed on Day 43 – representing proteins that remained after the washes – were dimers of DsRed. (No image available.)
On the second de-stained gel, surprisingly, there were no large unidentified bands present with our proteins (lanes 7-9):
Figure 34: The results of SDS-PAGE for our M3 Red elutions. Although the gel was not de-stained properly, bands could still be seen for the neat protein and the dilutions (to 1 in 3). (The angle at which the photograph was taken makes the bands appear further down the gel than they actually were.) No bands for the suspected dimers were visible on this gel.
We could see three large bands for our neat protein, our 1:2 dilution and our 1:3 dilution respectively, and virtually nothing else. Given that there had been no further washing steps since that time, along with the fact that contaminant proteins don’t just disappear, we had to conclude that the dimers separated; this may have occurred when our samples were left on the heat block.
The gel wasn’t de-stained properly, as we ran out of de-staining solution and the components to make it up (MEOH and glacial acetic acid). However, we could still see another small band for our 1:2 dilution on the gel. If this is an issue, then Jonathan and Donal still have the neat sample and the 1:3 dilution to work with.
Whatever the explanation may be, the results of this gel indicate that we did indeed obtain a pure solution of enhanced DsRed. This marked the end of the project.
The pragmatic goal of this project was to obtain an enhanced form of DsRed, such that its fusion with recombinant lectins enables the easier detection of early-stage apoptosis in CHO cells subjected to stress. Not only was this goal achieved – in the form of E5up with V71M, fluorescing magenta under UV light – but the ISSC may also now have a green-emitting version of DsRed, depending on the success of our final PCR. Using the ligated plasmid obtained on Day 46, when the time is right, Donal Moynihan and Jonathan Cawley can proceed with transformation to determine if this is the case.
The broader goal of the project was to become acquainted with a series of techniques that are useful for anyone pursuing a career in practical biology. This goal was achieved and then some: for not only did we learn how to carry out site-directed mutagenesis, transformation, protein expression analysis and IMAC, but we were also given the opportunity to refine existing techniques, such as streak plating, DNA purification and gel electrophoresis.
There was a more philosophical aspect to the project as well. We learned – with great difficulty – not to panic when results go awry, as well as to expect the unexpected. We were taught that making mistakes is necessary to fully understand and appreciate any technique, be it simple or complex. These are lessons that can only be learned with experience, not from a textbook. To provide a sample of the encouragement given to us by Dr. O’Connor:
“Kareem, any laboratory that claims to have a 80 percent success rate is lying; it’s probably around 20 percent.”
Given the frequency of our mistakes, hearing this on a regular basis was most reassuring.
I would like to offer my sincere thanks to Dr. Brendan O’Connor for his almost superhuman enthusiasm for glycosylation, and for his quick replies to my queries.
I must also thank Jonathan Cawley, Donal Moynihan and Disha Choudary for their constant assistance and sage advice, in spite of the mountain of work that rests upon their shoulders.
Lastly, I would like to thank Dr. Michael O’Connell and Dr. Anne-Parle McDermott for their constant support over the past four years, in good times and bad. I am certain that without their encouragement, I would never have progressed to fourth year at all.
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Appendix A: The amino acid sequence of DsRed
DsRed is 225 amino acids in length, or 675 base-pairs. This translates roughly into a protein that is 27.5 kDa in size. The poly-His tag is fused to the N-terminal of the protein. DsRed is located within the multiple cloning site of pQE-30.
Source: Terskikh et al. (2001), ‘Analysis of DsRed Mutants: Space Around the Fluorophore Accelerates Fluorescence Development.’ J. Biol. Chem. 2002, 277: 7633-7636.
Appendix B: The nucleotide sequences of our mutagenic primers
Each set of primers incorporates one SNP, to be induced in succession by mutagenesis PCR. There is a need for three sets of primers as the nucleotides to be altered in DsRed are not in close proximity to one another. A primer incorporating all three mutations would have to be at least 378 bp in length, which is obviously not feasible.
Appendix C: Table of the amino acids and their respective codons
This table represents the redundancy of the genetic code: that is, how an amino acid may be represented by more than one codon. Leucine, for example, is coded for by six different codons. This is advantageous, for if a mutation occurs during transcription making one of those six different codons non-functional, then the other five can still code for Leucine.
Appendix D: Diagram of the mutagenesis process and primer design