LABORATORY 9 CONSTRUCTION OF A FUSION PROTEIN CONSTRUCT TRANSFECTION OF MAMMALIAN CELLS AND VITAL STAINING OF ORGANELLES

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1 LABORATORY 9 CONSTRUCTION OF A FUSION PROTEIN CONSTRUCT TRANSFECTION OF MAMMALIAN CELLS AND VITAL STAINING OF ORGANELLES PURPOSE OF THE LABORATORY The purpose of this laboratory is to introduce you to a very common technique used in cell biology research: transfection of DNA into mammalian cells to study where the expressed protein is localized. We have made three plasmids for you, which we will call X, Y, and Z. Each plasmid contains a gene encoding a fluorescent fusion protein, in which the coding sequence for green fluorescent protein (GFP) from the jellyfish Aequorea victoria is fused to a segment of coding sequence for a protein that we want to investigate. Specifically, in this lab we hope to determine where the proteins of interest are targeted within the cell. The intrinsic fluorescence of GFP enables us to visualize the localization of the fusion proteins in living cells. During the previous lab (the cell cycle lab), we will also construct a fourth plasmid fusing a segment of a gene from C. elegans to GFP so that you can learn the techniques used for this type of work. You will introduce the plasmids into cultured human cervical carcinoma cells (HeLa cells) by DNA transfection. Over the next couple of days, the transfected cells should synthesize the GFP fusion proteins and target them to the appropriate location in the cell, based on the sorting information in sequences X, Y, or Z. To analyze the protein localization, we will observe the fluorescence from the GFP fusion proteins after staining the transfected cells with fluorescent vital stains, fluorescent molecules that can be used to stain living cells and that localize to known cellular compartments. These dyes have been chosen in part because they fluoresce at different wavelengths than GFP (either blue emission or red emission), so that you can distinguish between the vital stains and the GFP proteins by imaging them using different filters. This should enable you to identify the compartment of the cell where the GFP fusion protein is detected. I. TRANSGENE CONSTRUCTION AND DNA TRANSFECTION Introduction After identifying an intresting gene or protein through biochemical or genetic methods, a researcher will typically start to address the fundamental question, So, what does it do? An important tool for analysis of protein structure and function is the ability to introduce DNA transgenes (meaning any foreign gene that is introduced into a cell experimentally) into cultured cells or organisms. For example, introduction of transgenes can be used for any of the following experimental goals (and there are many others): 1

2 1) The intracellular targeting (or localization) of the protein can be examined by fluorescent detection in various cell types this is the primary application we will focus on in this laboratory. 2) The function of a protein on individual cells can be studied by analyzing the effects of expressing or overexpressing the protein from a transgenic construct. 3) A protein of interest can be expressed in cells at high levels and then purified for biochemical or structural studies. 4) The protein can be mutagenized by altering the coding sequence, and the effects of such alterations on the localization or function can be studied in a variety of ways. 5) The genetic elements responsible for controlling gene expression (where, when, and how abundantly the gene is transcribed and translated) can be studied by altering the regulatory information and then examining the resulting protein expression pattern. One of the first steps that we typically take in studying the function of a new protein is to determine where within the cell a protein of interest is localized. One way to do this is to raise an antibody against the purified protein and use it localize the protein by immunofluorescence, as you did in the cytoskeleton lab. An alternate approach is to use recombinant DNA methods to fuse the gene encoding the protein of interest to a gene encoding a fluorescent protein, introduce the resulting DNA construct into cultured cells, and visualize the resulting fluorescent fusion protein. This method is frequently faster than raising an antibody, since DNA constructs can be generated inexpensively in a matter of a few days (or less) and transfection is also a very rapid technique. By contrast, it usually requires at least 2 months and roughly $1000 to generate an antibody, and not all antibodies will ultimately work for immunofluorescence. Also, by expressing a protein of interest in cultured cells we can study its localization even if it is not normally expressed in that type of cell. For example, a protein that is normally expressed only in the mitochondria of liver cells will also be targeted to the mitochondria of HeLa cells (which are derived from an ovarian tumor) if the gene is expressed artificially in those cells. In this lab we will construct a plasmid by taking part of a particular gene sequence from one organism (the nematode worm Caenorhabditis elegans) and fusing it to a gene for the GFP protein from another organism, the jellyfish Aequorea victoria, to determine where this worm/jellyfish fusion protein localizes within human cells. If this experiment works (and we don t know yet, since we haven t done it before), it s because the genetic code (the code that cells use to convert DNA into RNA and then protein) and the protein sorting information that determines where individual proteins are targeted within cells are widely conserved. Production of a GFP fusion construct In this lab you will be given three different plasmids encoding GFP fusion proteins X, Y, and Z. In addition, we will use molecular biological techniques to construct a new plasmid that should express fusion protein U. To produce this plasmid, we will insert a piece of coding sequence from a gene from the nematode C. elegans, which was amplified by PCR from genomic DNA, into an appropriate cloning vector. The amplified coding sequence will be fused in frame to the coding sequence of jellyfish green fluorescent protein (GFP). Since you have previously 2

3 carried out both PCR and restriction digests, the staff has already done this part of the cloning procedure, and you will pick it up at the next step. Specifically, you will purify the linearized, digested vector DNA and the digested PCR product. You will then ligate these two pieces of DNA together to create a new plasmid, transform the ligated DNA into competent bacterial cells, select for the presence of the plasmid, and then purify the plasmid and verify that its structure is correct. If you successfully generate this plasmid, you can transfect it into tissue culture cells along with the other three plasmids we are providing for you. Transfection methods Transfection is the general term for techniques in which DNA is introduced into eukaryotic cells by chemical methods (there are other ways to get foreign DNA into cultured cells, including injection or infection with a specially engineered virus). A sequence of interest is typically inserted into an appropriate cloning vector (also called a plasmid), introduced into bacteria (introduction of foreign DNA into bacterial cells is usually called transformation), amplified by replication in the bacterial cells, and then purified from the bacteria. To transfect the DNA across the plasma membrane of mammalian cells requires that it be packaged using transfection reagents to allow its uptake by the cells. An alternative method is electroporation, which involves exposing the cells to a strong electrical field, which makes transient holes in the membranes and allows DNA to enter (1). Following transfection, the plasmid vector is designed to drive the expression of the transgene inside the cell. Plasmids can be transfected into mammalian by several different techniques, the most common of which are listed below. The optimal method for transfecting cells will vary based on the size of the transgenic construct and the particular cell types. To allow DNA, which has a strong negative charge, to cross the lipid membrane of living mammalian cells, the negative charged phosphate backbone must be neutralized, or countered, by positively charged ions. Commonly used transfection methods include the following: DEAE-dextran (2) The positively charged polymer DEAE-dextran binds to DNA because of its negatively charged phosphate backbone, and also sticks to the plasma membrane of cells. Cellular uptake occurs by endocytosis. Calcium phosphate precipitation (3) Positively charged calcium ions bind to and neutralize the negatively charged phosphate groups of the DNA. The addition of phosphate precipitates the Ca-DNA complexes, which attach to the cell and are endocytosed. Liposome-mediated transfection (4) Cationic (positively charged) lipids form small liposomes with a positively charged surface. These liposomes can be used to package DNA if the lipids are added to an appropriate quantity of DNA. Charge interactions with DNA phosphate groups and the plasma membrane mediate binding and fusion of the DNA/lipid complex to the cell. We will use this method in this laboratory. The mammalian cell transfection method we will use in this laboratory results in transient transfection, meaning that the DNA we introduce into the cell will stay 3

4 around for a while but would not permanently affect the cell culture, if we were to keep the cells growing for many generations. In transient transfections, the DNA is maintained extrachromasomally (meaning as a separate piece of DNA in the nucleus). Since the DNA does not replicate or attach to the mitotic spindle efficiently, it will be lost as the cells replicate and divide, typically over hours. Protein expression from the transfected DNA can start as soon as the DNA reaches the nucleus and is transcribed to make mrna. The level of protein expression is determined in part by the number of copies of DNA that reaches the nucleus of the cells, which is referred to as the transfection efficiency. Different cells in a culture will show variable levels of expression you will see this as bright GFP in some cells and none in others. In some experiments, it is important that each cell in a culture contain the same dose of a gene and/or that the gene be steadily expressed over a long time. For such work, researchers use a variation of the transfection method called stable transfection. This starts out identically to transient transfection, but a selectable marker such as a gene producing drug resistance (usually to neomycin) must be included in the vector. At a low frequency, the vector will integrate into one of the cell s normal chromosomes by a spontaneous recombination event. Although this does not happen in many cells, we can kill off all cells except for those few that have stably integrated the transfected DNA by exposing cells to the drug. These resistant cells are then allowed to multiply to generate a stably transfected cell line. In addition to choosing between transient and stable transfection methods, in designing the expression vector a researcher must decide what kind of promoter will drive the expression of the transgene in the eukaryotic cells. If the protein is not toxic to cells, it may be fine to use a constitutive promoter (one that is turned on all the time). In this lab, our transgenes are driven by pcmv, which is a very strong constitutive promoter (i.e., this promoter sequence drives high transcription of the gene all the time). Sometimes a protein will be toxic to cells or there will be some other experimental reason to turn the gene on only at a specific time. For such experiments, an inducible promoter is used. These promoters are usually off but can be turned on by changing the environmental conditions for example, by heat shocking the cells or by adding a drug such as tetracycline to the growth medium. II. PROTEIN SORTING AND VITAL STAINING OF ORGANELLES Introduction All proteins in the cell translated, or synthesized from mrna, by ribosomes. Most ribosomes are in the cytosol, either floating around or anchored to the membrane of the Endoplasmic Reticulum. The ultimate cellular destination of the proteins is determined by signals within their amino acid sequences. Proteins that do not have a sorting signal remain in the cytosol by default. Others have specific sorting signals that direct their transport from the cytosol into the nucleus, the ER, the mitochondria, plastids (in plants), or peroxisomes. Sorting signals can also direct the transport of proteins from the ER to other destinations in the cell. There are three different ways by which proteins move from one compartment to another: 1) Gated transport. Transport between the nucleus and cytoplasm occurs by this mechanism. The protein traffic occurs between topologically equivalent spaces, which are connected by nuclear pore complexes, which allow free diffusion of 4

5 small molecules but actively transport specific proteins and macromolecular assemblies. 2) Transmembrane transport. The initial translocation of proteins from the cytosol into the ER or the mitochondrial matrix occurs by this mechanism. Membranebound protein translocators directly transport specific proteins across a membrane from the cytosol into a space that is topologically distinct. The protein molecule usually must unfold to be snaked through the membrane. 3) Vesicular transport. Transport between the ER and Golgi, and secretory and endocytotic trafficking occur as transport vesicles ferry proteins from one compartment to another. Signal Sequence and Signal Patch The sorting signals that target proteins to individual compartments can be in the form of either signal sequences or signal patches. A signal sequence is a contiguous stretch of amino acid sequence about residues long. It is often removed from the polypeptide once the protein reaches its target compartment. Some typical signal peptides are shown below. Positively charged amino acids are in bold, negatively charged amino acids are in bold italic. Stretches of hydrophobic amino acids are outlined Function of Signal Peptide Import into ER Retention in lumen of ER Import into mitochondria Import into nucleus Example of Signal Peptide H 2 N-M-M-S-F-V-S-L-L-L-V-G-I-L-F-W-A-T-E-A-E- Q-L-T-K-C-E-V-F-Q- -K-D-E-L-COOH H 2 N-M-L-S-L-R-Q-S-I-R-F-F-K-P-A-A-T-R-T-L-C-S- S-R-Y-L-L P-P-K-K-K-R-K-V By contrast, a signal patch consists of a specific three-dimensional structure on the protein s surface; this structure can be formed from discontinuous regions of a polypeptide chain after it is folded into the native conformation. As a consequence, signal patches are much more difficult to identify than signal sequences. It is often difficult to deduce a protein s cellular location by simple inspection of its primary structure. For this reason, a common first step to investigate the function of an unknown protein is to transfect its coding cdna into cells and determine the protein s cellular location experimentally. This is the approach you will take in this week s exercise. Expression of GFP Fusion Proteins The cdna encoding the unknown proteins you will be transfecting has been fused to the green fluorescent protein (GFP) sequence. GFP is a naturally fluorescent protein isolated from the jelly fish Aequorea victoria. The protein is not fluorescent immediately after synthesis, but becomes fluorescent within an hour after a selfcatalyzed post-translational modification. Proteins tagged with the GFP sequence 5

6 can be visualized by fluorescence microscopy in living cells. This property has made it a popular reporter protein for cell biological research. Derivatives of GFP, such as EGFP (enhanced green fluorescent protein), YFP (yellow fluorescent protein), and CFP (cyan fluorescent protein) have also been engineered and are widely used. More recently, a red fluorescent protein from a reef coral (DsRed) has also been harnessed and re-engineered for experimental use. To express a GFP-tagged protein, the DNA sequence encoding GFP is fused to the gene for the protein of interest either at the C-terminus or N-terminus, as shown below for Protein X and Protein Y, respectively. The GFP sequence can also been inserted in the middle of an unknown protein, as we have done for Protein Z (see below). This fusion construct is then inserted into a mammalian expression vector. The final plasmid is amplified and used for DNA transfection. You should understand the function of the different elements in the cloning vector: P CMV : Cytomegalovirus Immediate Early Promoter, drives transgene expression at high levels in mammalian cells. GFP: Green Fluorescent Protein Open Reading Frame (ORF). BGH pa: Bovine Growth Hormone polyadenylation sequence f1 : bacterial origin of replication that allows rescue of single-stranded DNA SV40 (Arrow): Simian Virus 40 Early Promoter; drives expression of the Neomycin resistance gene in mammalian cells. Neomycin: Drug resistance gene for selection of stable transformants of mammalian cells. SV40 (Box): Polyadenylation signal for Neomycin gene, derived from Simian Virus 40. Ampicillin: Drug resistance gene with bacterial promoter for expression in E. coli puc: Bacterial origin of replication, allows propagation and amplification of plasmid in E. coli. 6

7 Vital Staining of Organelles Intracellular membranes of different organelles in eukaryotic cells are quite varied in both their lipid and protein components. Moreover, the size and shape of individual organelles is strongly dependent on the cell type studied. It is often difficult to determine the identity of an organelle based only on its particular appearance in a particular cell type. To identify organelles and other subcellular compartments, cell biologists often employ special vital stains that are reliably targeted to individual organelles due to their biochemical properties. In this lab, we are attempting to identify the cellular compartment where several different fluorescent fusion proteins are targeted. To achieve this goal, we will use a double-staining method in which the distribution of each unknown protein is directly compared to the staining pattern of a known organelle marker within the same cell. A protein is judged to be in a given organelle only if its pattern of distribution matches that of the known organelle marker. Keep in mind that the level of resolution of light microscopy is sometimes insufficient to resolve individual structures. Therefore, different organelles may produce partially overlapping patterns. The stains used in this lab are called vital stains because they are taken up and localized within living cells. One of the primary advantages of vital staining is that the dynamics of the stained structure can be examined in the living cell. (In contrast, the fluorescence staining methods you used in the cytoskeleton lab were performed on fixed [dead] cells.) We will use the following five vital stains: (1) Golgi vital stain: BODIPY-TR ceramide. Ceramide is a lipid that consists of sphingosine, an amino-alcohol with a long hydrocarbon chain, that is amide bonded to a fatty acyl chain. Within cells, ceramide is metabolized within the lumenal leaflet of Golgi membranes by the addition of polar head groups to generate sphingomyelin and glycolipids. You will be labeling cells by addition of BODIPY-TR ceramide, a modified version of ceramide in which the acyl chain has conjugated to a fluorphore, BODIPY- TR, which has an emission maximum arount 617 nm, similar to rhodamine (Invitrogen Molecular Probes Cat. No. D-7540). In the absence of metabolism (for example at 4 C), the fluorescence will be distributed throughout the cell. However, upon warm-up the normal metabolism of the BODIPY-TR ceramide (addition of head groups in the Golgi complex) prevents it from flipping into the cytoplasmic leaflet and as a consequence the fluorescence becomes trapped in the Golgi complex (5). The BODIPY-TR ceramide is complexed to the protein BSA (bovine serum albumin), which makes the lipid soluble in aqueous buffer such as PBS or tissue culture medium. (2) Endosome vital stain: Rhodamine-transferrin. Cells internalize a variety of extracellular molecules by receptor-mediated endocytosis. Here we will visualize the endosomes by adding a fluorescently labeled protein, transferrin, which can be taken up by living cells and concentrated in the endosome. The normal function of transferrin is to transport iron in the blood so that it can be delivered to cells that 7

8 produce heme-containing proteins such as hemoglobin (produced in immature red blood cells). Transferrin receptors on the plasma membrane bind to transferrin which is bound to iron (ferrotransferrin) and become incorporated into clathrincoated endocytic vesicles. These vesicles deliver their cargo to the early endosome, where the low ph causes the iron to dissociate from transferrin. The iron-free transferrin (apotransferrin), still bound to the transferrin receptor then recycles back to the cell surface via recycling endosomes. Once delivered to the plasma membrane, the iron-free transferrin dissociates from the receptor and is free to bind more iron (6). You will be incubating cells with rhodamine-labeled transferrin in order to follow this pathway in living cells. (3) Mitochondria vital stain: MitoTracker Red. This is a stain developed by Molecular Probes (MitoTracker Red CM-H2XROS, Cat. No. M-7513). The unmodified dye is membrane-permeant (meaning it goes through lipid membranes) and non-fluorescent. It passively diffuses through the plasma membrane and throughout the cell. Upon oxidation in the special environment of the living mitochondria, the dye becomes fluorescent, and also becomes permanently trapped inside the mitochondria. If desired, after incubation with MitoTracker Red, cells can be fixed and stained with other markers the fluorescence remains in the mitochondria. (4) Endoplasmic Reticulum vital stain: ER-Tracker Blue-White. This stain was also developed by Invitrogen Molecular Probes (ER-Tracker Blue-white DPX, Cat # E-12353). For reasons that are not fully understood, this that is selective for endoplasmic reticulum (ER) in live cells. It is non toxic and does not stain mitochondria. 5) DNA/Nuclear vital stain: Hoechst 33258/ You have already used one of these DNA-specific dyes in the apoptosis lab. Fluorescent DNA dyes such as DAPI and Hoechst have an advantage in that they become much more fluorescent upon binding to the minor groove of DNA, especially to AT-rich sequences. Therefore, if the dyes are used at an appropriate concentration (typically 0.2 to 5 μg/ml), the unbound dye does not result in high background. The Hoechst dyes are more cellpermeant than DAPI, and are thus more useful as vital stains. For optimal labeling of living cells, an incubation time of minutes is recommended. Because Hoechst dyes fluoresce strongly in the blue portion of the spectrum, they can be used together with GFP and any of the other vital stains in this lab except for ER-Tracker Blue-White. REFERENCES 1) Neumann, E., Schaefer-Ridder, M., Wang, Y., and P.H. Hofschneider (1982) Gene transfer into mouse lyoma cells by electroporation in high electric fields. EMBO J. 1: ) Vaheri, A. and J.S. Pagano (1965) Infectious poliovirus RNA: A sensitive method of assay. Virology 27: ) Graham, F.L. and A.J. Van der Eb. (1973) A new assay for the infectivity of human adenovirus 5 DNA. Virology 52:

9 4) Felgner, P.L., et al. (1987) Lipofection: A highly efficient, lipid-mediated DNA transfection procedure. PNAS 84: ) Lipsky NG; Pagano RE. (1985) A vital stain for the Golgi apparatus. Science 228:745. 6) Presley JF; Mayor S; Dunn KW; Johnson LS; McGraw TE; Maxfield FR (1993) The End2 mutation in CHO cells slows the exit of transferrin receptors from the recycling compartment but bulk membrane recycling is unaffected. J. Cell Biol. 122:

10 DETAILED INSTRUCTIONS Part A. Synthesis of a plasmid encoding a novel fusion protein (This will be carried out over three lab periods, two during the week of November while we do the cell cycle lab, and then the additional lab period just before Thanksgiving.) Each group will be given a tube containing 2 μg of vector pcmv/gfp/mcs and a separate tube containing 2 μg of the digested insert, T05G5.9_Cterm. This is a 240-bp double-stranded piece of DNA containing a portion of coding sequence from the Caenorhabditis elegans gene T05G5.9. You will need to gel-purify both the vector and the insert to separate these fragments from the small pieces that were cleaved off by the Not I and Xba I restriction enzymes. Because the vector is much larger than the insert, it is best to do these two purifications using two different agarose gels. The table below shows the typical range of fragment sizes that gels of different agarose concentration are useful for resolving (adapted from Molecular Cloning, Third Edition, edited by Sambrook and Russell). Agarose concentration in gel (% w/v) Range of separation of linear DNA fragments (kb) You should use a 2% (w/v) gel to purify the small insert DNA, which is only 240 bp (0.24 kb) long, and a 0.8% gel to purify the linearized vector, which is 5.8 kb long. You will find agarose prepared for you in 50-ml conical tubes in the 70 C waterbath. Add 2.5 μl of 10 mg/ml EtBr solution (ethidium bromide) before pouring each gel. You should share one of each of these gels with the three other groups at your same bench load the fragments leaving a blank lane in between, and run the gel after all groups have loaded their DNA. The small insert fragment will run slightly slower than the purple dye in the loading buffer, so you should run the gel until the dye front is about halfway down or further. You can run the gel to purify the 5.8-kb vector fragment for about the same amount of time, or a bit longer (anything else on the gel will be very small and easily resolved from this fragment. To purify the fragments: we will use Qiagen gel extraction kits. Each pair of partners will receive two kits, one for the insert and one for the vector. The manual is available in the teaching lab, and can also be downloaded as a PDF online at tion/qq_spin/ _qiag_complete.pdf 10

11 You will be following the procedure for QIAquick Gel Extraction using a microcentrifuge, which is on p of the manual. Ethanol will be added to Buffer PE for the entire class. Note: since everyone will be using the same reagents for gel binding, column washing, and eluting their DNA, you should be very careful not to contaminate these solutions. Use only clean pipet tips to remove what you need from the common bottles of solution. You will start by cutting your DNA out of of the preparative agarose gels. You will visualize the DNA bands by the fluorescence of ethidium bromide using a UV transilluminator. Note that you should work quickly to minimize exposure of the DNA to the UV light, which can damage it, resulting in mutations or failure of the cloning. Make 4 cuts around the desired band with a sharp razor blade to excise the DNA fragment in the minimum possible volume of agarose. Using forceps, transfer the small chunk of agarose containing your DNA to a preweighed 1.5-ml tube (note the weight of the empty tube in your notebook). Then weigh the tube again to calculate the volume of the agarose chunk and follow the procedure for QIAquick Gel Extraction using a microcentrifuge from the Qiagen handbook. This basic protocol is included in the next two pages of this manual, but you may want to consult the more extensive information in the manual to learn more about how this method works, how much DNA you can expect to recover, and potential problems you might encounter. The DNA will be eluted from the purification columns in a volume of 50 μl. If you recovered 100% of your DNA fragments, this would give you a concentration of 2 μg/50 μl, or 40 μl/ml. A more realistic estimate is that you will recover about 50% of the DNA in each band. You can determine the exact concentration of your recovered DNA using the Nanodrop spectrophotometer. Knowing precisley the concentration of DNA in your tube is a good practice in the lab, and the Nanodrop makes it easy to do this with minimal loss or dilution of your sample. 11

12 QIAquick Gel Extraction Kit Protocol using a microcentrifuge This protocol is designed to extract and purify DNA of 70 bp to 10 kb from standard or low-melt agarose gels in TAE or TBE buffer. Up to 400 mg agarose can be processed per spin column. This kit can also be used for DNA cleanup from enzymatic reactions (see page 8). For DNA cleanup from enzymatic reactions using this protocol, add 3 volumes of Buffer QG and 1 volume of isopropanol to the reaction, mix, and proceed with step 6 of the protocol. Alternatively, use the MinElute Reaction Cleanup Kit. Important points before starting The yellow color of Buffer QG indicates a ph 7.5. Add ethanol (96 100%) to Buffer PE before use (see bottle label for volume). All centrifugation steps are carried out at 10,000 x g in a conventional table-top microcentrifuge at room temperature. Procedure 1. Excise the DNA fragment from the agarose gel with a clean, sharp scalpel. Minimize the size of the gel slice by removing extra agarose. 2. Weigh the gel slice in a colorless tube. Add 3 volumes of Buffer QG to 1 volume of gel (100 mg ~ 100 µl). For example, add 300 µl of Buffer QG to each 100 mg of gel. For >2% agarose gels, add 6 volumes of Buffer QG. The maximum amount of gel slice per QIAquick column is 400 mg; for gel slices >400 mg use more than one QIAquick column. 3. Incubate at 50 C for 10 min (or until the gel slice has completely dissolved). To help dissolve gel, mix by vortexing the tube every 2 3 min during the incubation. IMPORTANT: Solubilize agarose completely. For >2% gels, increase incubation time. 4. After the gel slice has dissolved completely, check that the color of the mixture is yellow (similar to Buffer QG without dissolved agarose). If the color of the mixture is orange or violet, add 10 µl of 3 M sodium acetate, ph 5.0, and mix. The color of the mixture will turn to yellow. The adsorption of DNA to the QIAquick membrane is efficient only at ph 7.5. Buffer QG contains a ph indicator which is yellow at ph 7.5 and orange or violet at higher ph, allowing easy determination of the optimal ph for DNA binding. 5. Add 1 gel volume of isopropanol to the sample and mix. For example, if the agarose gel slice is 100 mg, add 100 µl isopropanol. This step increases the yield of DNA fragments <500 bp and >4 kb. For DNA fragments between 500 bp and 4 kb, addition of isopropanol has no effect on yield. Do not centrifuge the sample at this stage. Gel Extraction Spin Protocol QIAquick Spin Handbook 03/

13 Gel Extraction Spin Protocol 6. Place a QIAquick spin column in a provided 2 ml collection tube. 7. To bind DNA, apply the sample to the QIAquick column, and centrifuge for 1 min. The maximum volume of the column reservoir is 800 µl. For sample volumes of more than 800 µl, simply load and spin again. 8. Discard flow-through and place QIAquick column back in the same collection tube. Collection tubes are reused to reduce plastic waste. 9. Recommended: Add 0.5 ml of Buffer QG to QIAquick column and centrifuge for 1 min. This step will remove all traces of agarose. It is only required when the DNA will subsequently be used for direct sequencing, in vitro transcription, or microinjection. 10. To wash, add 0.75 ml of Buffer PE to QIAquick column and centrifuge for 1 min. Note: If the DNA will be used for salt-sensitive applications, such as blunt-end ligation and direct sequencing, let the column stand 2 5 min after addition of Buffer PE, before centrifuging. 11. Discard the flow-through and centrifuge the QIAquick column for an additional 1 min at 10,000 x g. IMPORTANT: Residual ethanol from Buffer PE will not be completely removed unless the flow-through is discarded before this additional centrifugation. 12. Place QIAquick column into a clean 1.5 ml microcentrifuge tube. 13. To elute DNA, add 50 µl of Buffer EB (10 mm Tris Cl, ph 8.5) or water to the center of the QIAquick membrane and centrifuge the column for 1 min. Alternatively, for increased DNA concentration, add 30 µl elution buffer to the center of the QIAquick membrane, let the column stand for 1 min, and then centrifuge for 1 min. IMPORTANT: Ensure that the elution buffer is dispensed directly onto the QIAquick membrane for complete elution of bound DNA. The average eluate volume is 48 µl from 50 µl elution buffer volume, and 28 µl from 30 µl. Elution efficiency is dependent on ph. The maximum elution efficiency is achieved between ph 7.0 and 8.5. When using water, make sure that the ph value is within this range, and store DNA at 20 C as DNA may degrade in the absence of a buffering agent. The purified DNA can also be eluted in TE (10 mm Tris Cl, 1 mm EDTA, ph 8.0), but the EDTA may inhibit subsequent enzymatic reactions. 14. If the purified DNA is to be analyzed on a gel, add 1 volume of Loading Dye to 5 volumes of purified DNA. Mix the solution by pipetting up and down before loading the gel. Loading dye contains 3 marker dyes (bromophenol blue, xylene cyanol, and orange G) that facilitate estimation of DNA migration distance and optimization of agarose gel run time. Refer to Table 2 (page 15) to identify the dyes according to migration distance and agarose gel percentage and type. 26 QIAquick Spin Handbook 03/2006

14 Set up your ligations. Typically, ligations are performed with about a 3:1 molar ratio of insert:vector DNA. You can measure the concentration of the DNA you purified from the gel using the Nanodrop spectrophotometer to calculate precisely how much you need to mix from each tube to get a 3:1 molar ratio. Alternatively, assume that the vector and insert DNA were recovered with equal efficiency from the gels and figure out the relative volume of each tube you need to mix. Your total volume of [insert + vector] should be 8 μl. Notes: The ratio of insert to vector in the ligation is more important than the total quantity of DNA. Since the insert DNA is much smaller than the vector, you need much less by weight about 8-fold lower amount of the insert by weight will result in 3-fold higher molar concentration of the insert. In addition to carrying out a ligation of the [vector + insert DNA], you should do a minus insert (vector alone) control using the same amount of vector DNA but replacing the volume of the insert with sterile water. This will give you some idea of your background how many of the transformed colonies you see after your ligation and transformation are likely to be what you want that is, the insert ligated into the vector. The total volume of each ligation should be 10 μl. Set up the reactions on ice. (X) μl volume of vector (8 X) μl volume of insert (or water for control ligation) 1.0 μl 10X buffer for T4 DNA ligase 1.0 μl T4 DNA ligase enzyme 10 μl total volume Mix the components of the reaction by gently tapping the tube. When your ligation is ready to go, place it in a rack on your benchtop (~22 C) for one hour. Transformation After your ligations have been incubating for 1 hour, you should transform the DNA into competent bacterial cells. You will need one tube of competent cells for the [vector + insert] ligation and one tube for the vector alone control. Competent cells should be kept ON ICE. These cells have been specially prepared to allow them to take up exogenous DNA by transformation. An alternative method to get DNA into bacterial cells is electrotransformation, which requires a special device called an electroporator, which exposes the cells to an electrical field. 1. Label the bottoms of two LB-amp plates with your name, your partner s name, and your GSI s name. 2. Label one plate vector only and one plate vector + insert 3. Chill your ligation mix on ice for 5 minutes 4. Pipet 3 μl of cold ligation mixture into one tube of competent cells (30 μl). 5. Mix the DNA with the bacteria gently by tapping the tube for a few seconds. 6. Return the tube to ice and incubate for 30 minutes. 7. Add 0.5 ml of LB or SOC medium 12

15 8. Cap tube and incubate cells for 30 minutes in a 37 C waterbath. 9. Using a clean, sterile pipet tip, transfer 200 μl of cells to an LB-Ampicillin plate. Using a sterile glass spreader, spread the cells evenly around the surface of the plate (try not to break the surface of the agar. ). Cover the plate with the lid. 10. Once the liquid has dried, invert the plates so that the cover is on the bottom. This minimizes the condensation of water on the lid, which can drip onto the agar and cross-contaminate your colonies. 11. Give your plates to your GSI, who will place them in an incubator until the next class. Next lab period (November 15/16): Assess the success of the ligation/ transformation and pick single colonies to prepare DNA minipreps Count the total number of colonies on the vector alone and vector + insert plates. Ideally you will have lots of colonies (>50) on the [vector + insert] plate and few or none on the control plate. If there are a huge number (>>100) on the vector + insert plate, you don t need to count them all; try to divide the plate into 8 equal sectors by drawing a pizza-slice pattern on the bottom or top of the petri plate with a sharpie, then count the colonies in 1 sector and multiply by 8 to estimate the total number on the plate. Record the number of colonies on each plate in your notebook. Sometime during the lab period, you and your partner should pipet 2 ml of LBamp (provided) into each of four sterile culture tubes, label the tubes with your names and sample numbers (1-4), and then use a sterile toothpick to transfer bacteria from a single bacterial colony from the [vector + insert] plate into each of the tubes containing liquid medium. Your GSI can help show you how to do this. These cultures will be grown up before the next lab period, which will be during the following week. If you do not have any colonies on your [vector + insert] plate, see if you can get some from another group that has had better luck with their ligation and transformation. You are picking four colonies (rather than just one) to try to ensure that at least one of the transformants has the correct vector, which you will test by miniprepping the DNA and carrying out a diagnostic digest next week. If you had low background on the [vector alone] plate, there is a good chance that all 4 colonies will contain the correct vector. Next lab period (Monday/Tuesday before Thanksgiving): DNA minipreps and diagnostic digests of individual colonies from the transformation Collect your four bacterial cultures from the shaker or rack. They have grown for at least 8 hours at 37 C, and they should be highly turbid (cloudy) due to growth of the bacteria to saturation. You and your partner will get 4 QIAgen miniprep columns. You should carry out a separate miniprep to isolate the plasmid DNA from each bacterial culture using one QIAprep miniprep column, following the manufacturer s instructions closely. The manual is available in the lab and you can download it at 13

16 The protocol you need is on pp of the Handbook, but you may also find the information elsewhere in the manual to be useful. If your minipreps fail you may want to consult the troubleshooting guide starting on p. 36. Once you have completed the miniprep procedure, you should measure the DNA concentration using the Nanodrop spectrophotometer. This will be important for the subsequent digestion and transfection steps. You should then carry out a restriction digest of 1 μg of each plasmid miniprep using the enzyme Pst I. Because the vector contains a single Pst I site and the insert we ligated into the vector also contains a Pst I stie, this enzyme will cut the vector into two bands of approximately 5 kb and 0.8 kb. If the insert is not present, there will be a single band of about 5.8 kb due to the Pst I site in the vector. Based on your measured DNA concentration, calculate the volume of plasmid you will need from each miniprep to have 1 μg. Record these volumes in your notebook as X 1, X 2, X 3, and X 4. Set up four restriction digests in separate labeled 1.5-ml eppendorf tubes on ice: 16-X μl H2O X μl miniprep DNA 2 μl New England Biolabs Buffer 3 (10X stock) 1 μl 2 mg/ml BSA solution (100 μg/ml final; this helps to stabilize the Pst I enzyme) 1 μl (5 units) Pst I enzyme 20 μl total volume Cap the tubes and mix the restriction digest mixes gently by tapping, then place in a 37 C waterbath for 1 hour. While the DNA is digesting, you should pour a 1% agarose gel using the agarose provided, after adding ethidium bromide solution. Add 4 μl of 6X DNA loading buffer to the digested DNA and mix. Load half of each reaction (12 μl, including the DNA loading buffer) onto the gel. Include appropriate molecular weight markers. Run the gel for 30 minutes. Photograph the bands on the gel using the UV light box and polaroid camera. Tape the polaroid into your notebook and carefully label it in a way that you will remember what each sample was, and what the markers are. If one or more of your plasmids shows the correct digest pattern, you can use it in next week s transfection lab. You will need to know the concentration of the plasmid, which you measured before doing the restriction digest. 14

17 Part B. Transfection and Vital Staining (week of November 27-30) Day 1: Transfection of HeLa Cells The complete data set for this lab will include the five stains listed above for each of four unknown GFP construct transfections (X, Y, Z, and U) plus an unstained control. Each group will transfect four coverslips of HeLa cells with a DNA construct (Plasmid X, Y and Z, and U). The transfection method you will perform requires that the cells be seeded ~5 hours prior to adding the DNA and transfection reagents. In order to allow this lab to be completed in two days, the cells have been plated for you in advance (Part A). Proceed to Part B to start today s experiment. A. Seed cells on coverslips (5 hours prior to transfection): 1. Aspirate media from cells grown in 10-cm dishes. 2. Rinse each plate with 3 ml of Ca ++ /Mg ++ free PBS. Aspirate PBS. 3. Add 2 ml of Trypsin solution. Wait for 2-3 minutes to allow cells to detach. 4. Meanwhile, prepare a tube containing ~3 ml of fresh growth media. 5. Check cells on inverted scope, tapping dish to help dislodge cells. 6. Collect cells and transfer to tube with fresh growth media. 7. Centrifuge for 5 minutes at 1500 rpm. 8. Aspirate all but ~1 ml of supernatant, being careful not to disturb the cell pellets. 9. Flick to re-suspend cells and add media to bring total volume to 5 ml. 10. Count 10 μl on a hemocytometer. 11. Adjust volume to get 5 x 10 5 cells per ml. 12. Fill each well of a 6 well plate with 2 ml fresh growth media. 13. Add 500μl of cell suspension to each well and return the plate to incubator. (The above procedure was done for you at 9:30 am this morning.) 15

18 B. Transfect cells: You will be given three tubes containing different plasmids (X, Y and Z) each at a concentration of 40 ng/μl (TE buffer, ph 7-8) plus the reagents required for liposomemediated transfection. You should also measure the concentration of Plasmid U (the plasmid you constructed in the first part of this lab) using the Nanodrop spectrophotometer, if you haven t done so already and based on this measurement, calculate the volume you will need to transfer to a new tube to have 0.4 μg (400 ng). A list of reagents you will need is provided at the end of this section. You will be transfecting 4 coverslips, one with each plasmid. In 4 separate microfuge tubes prepare the following mixture for each coverslip. 1. Pipette 0.4 μg of your DNA sample into an 1.5 ml microfuge tube and add EC buffer to a final volume of 100μl. These reagents will be at your bench μl DNA + μl EC buffer=100μl final volume Repeat for second tube. 2. Add 3.2μl of Enhancer to each tube. Vortex briefly to mix. 3. Incubate at room temperature (RT) for 4 minutes. Spin down briefly in the tabletop centrifuge to collect any droplets on the walls or cap of the tube. 4. Add 2.5 μl of Effectene to each tube and mix by carefully pipetting up and down 5 times with a P-200 set to ~80 μl. Be careful not to introduce bubbles as the increased surface area leads to a higher oxygen exposure which can degrade some of the reagents. Do not depress the pipette beyond the first stop until the last cycle. 5. Incubate at RT for 5-10 minutes. 6. Obtain a 6 well plate containing three coverslips of HeLa cells from the incubator. Normally the following steps would be performed in a sterile hood, but with certain precautions it is not absolutely necessary. The growth media and PBS you will use are sterile. To aspirate the media and PBS, use a sterile plastic pipette and be extra careful not to touch it to any non-sterile surface. * 7. Aspirate media from wells. Rinse each well with 2 ml of sterile PBS and aspirate. 8. Add 1.6 ml of fresh growth media to each well. 9. With the P-1000 pipettor add 600μl of fresh media to one of the tubes containing DNA complex and mix by pipetting up and down twice. Immediately with the same pipette and tip add the mixture drop wise to one of the wells. Change tips and repeat for the second tube. Label the wells carefully and swirl gently to distribute. 10. Return your plate to the incubator. *Note: The media contains antibiotics that should suppress minor bacterial contamination for the two days before you stain and view the slide. Molds and fungi grow more slowly and should not be a concern within 48 hours. 16

19 Reagents List Growth Media: Dulbecco s Modified Eagle Medium (DMEM) + Fetal Calf Serum + Penicillin & Streptomycin PBS: 137 mm NaCl, 2.7 mm KCl, 4.3 mm Na 2 HPO 4, 1.47 mm KH 2 PO 4, ph 7.4 TE: 10 mm Tris, 1 mm EDTA, ph 7-8 adjusted with HCl EC-Buffer (Qiagen): Proprietary Content, salt buffer that facilitates DNA condensation with the enhancer. Enhancer (Qiagen): Proprietary Content, reagent that facilitates DNA condensation with EC-buffer. Effectene Reagent (Qiagen): Proprietary Content, lipid reagent that binds to condensed DNA. Day 2 Your group should have 4 transfected coverslips from the previous lab. To determine the subcellular localization of your unknown proteins, you will be counterstaining the cells with one of the vital dyes described in the introduction. Each group will perform one counter-stain such that every table of 4 groups (i.e. A&B, C&D and E&F) will have a complete set. The protocols generally vary in time such that you should not be crowded at the microscopes. The protocol for Mitotracker-Red and Blue-White ER are the closest to each other. The groups running the ER protocol should wait a few minutes after the Mitotracker group has started to begin their protocol. You will need to share your data to get the full set of experiments. Do not leave class until you have had the opportunity to observe each stain. All of these dyes are sensitive to light and should be kept in the dark as much as possible. Please read through all four protocols so that you have a good understanding of each. At the end of the four protocols are some general instructions pertinent to all groups. Check calculations with your GSI before proceeding with dilutions. 17

20 (A1, C1 & E1): Vital staining with MitoTracker-Red (Leave coverslips in the 6 well plate for this procedure) MitoTracker-Red is stored at a much higher concentration than it is added to the cells. The stock solution you are given is 10 μm. Make certain that your dilution is correct before proceeding. You can easily ruin your results by either having too much or too little of the compound. Remember that for this staining to work the mitochondria must be in good condition. Always keep in mind that the cells are alive and treat them with due care! 1. Make 2 ml of a 100 nm working solution of dye in growth media. μl Stock solution + μl growth media = 2ml working solution. (Optional: add 1 μl of 1 mg/ml Hoechst dye to the working solution to get a final concentration of 0.5 μg/ml) 2. Aspirate media from wells and rinse each well with ~2 ml PBS. 3. Aspirate PBS. Add 1 ml of working concentration dye in growth media to each well. 4. Incubate for 15 min at 37º C. 5. Prepare a glass slide with two 7 μl drops of Phenol-Red free media. 6. Aspirate dye containing media from wells and rinse each well with ~2 ml PBS. 7. Remove coverslips from the 6 well plate with forceps, wicking off excess PBS. Mount on the prepared glass slide. Seal with nail polish. View slides as soon as the nail polish is dry. (A2, C2 & E2): Vital staining with Blue-White-ER (Leave coverslips in the 6 well plate for this procedure) Blue-White_ER is stored at a much higher concentration than it is added to the cells. The stock solution you are given is 10 μm. Make certain that your dilution is correct before proceeding. You can easily ruin your results by either having too much or too little of the compound. 1. Make 2 ml of a 100 nm working solution of dye in growth media. μl Stock solution + μl growth media = 2ml working solution. 2. Aspirate media from wells and rinse each well with ~2 ml PBS. 3. Aspirate PBS. Add 1 ml of working solution of dye in growth media to each well. 4. Incubate for 30 min at 37º C. 5. Prepare a glass slide with two 7 μl drops of Phenol-Red free media. 6. Aspirate dye containing media from wells and rinse each well with ~2 ml PBS. 7. Remove coverslips from the 6 well plate with forceps, wicking off excess PBS. Mount on the prepared glass slide. Seal with nail polish. View slides as soon as the nail polish is dry. 18