Download 1 laboratory 9 construction of a fusion protein

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):
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1)
2)
3)
4)
5)
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.
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.
A protein of interest can be expressed in cells at high levels and then purified for
biochemical or structural studies.
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.
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
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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
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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 12-80 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
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2)
3)
small molecules but actively transport specific proteins and macromolecular
assemblies.
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.
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 15-60 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
Example of Signal Peptide
Import into ER
H2N-M-M-S-F-V-S-L-L-L-V-G-I-L-F-W-A-T-E-A-EQ-L-T-K-C-E-V-F-Q-
Retention in lumen of ER
-K-D-E-L-COOH
Import into mitochondria
H2N-M-L-S-L-R-Q-S-I-R-F-F-K-P-A-A-T-R-T-L-C-SS-R-Y-L-L
Import into nucleus
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
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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:
PCMV: 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.
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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, BODIPYTR, 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
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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 33258/33342. 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 20-30 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)
2)
3)
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: 841.
Vaheri, A. and J.S. Pagano (1965) Infectious poliovirus RNA: A sensitive
method of assay. Virology 27: 434.
Graham, F.L. and A.J. Van der Eb. (1973) A new assay for the infectivity of
human adenovirus 5 DNA. Virology 52: 456.
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4)
5)
6)
Felgner, P.L., et al. (1987) Lipofection: A highly efficient, lipid-mediated DNA
transfection procedure. PNAS 84: 7413.
Lipsky NG; Pagano RE. (1985) A vital stain for the Golgi apparatus. Science
228:745.
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.
122:1231.
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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
13-17 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)
0.3
0.6
0.7
0.9
1.2
1.5
2.0
Range of separation of linear
DNA fragments (kb)
5-60
1-20
0.8-10
0.5-7
0.4-6
0.2-3
0.1-2
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
http://www1.qiagen.com/literature/handbooks/PDF/DNACleanupAndConcentra
tion/QQ_Spin/1036019_Qiag_Complete.pdf
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You will be following the procedure for QIAquick Gel Extraction using a
microcentrifuge, which is on p. 25-26 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.
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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
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.
QIAquick Spin Handbook 03/2006
25
Gel Extraction
Spin Protocol
1.
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.
Gel Extraction
Spin Protocol
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
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
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
http://www1.qiagen.com/HB/QIAprepMiniprep
13
The protocol you need is on pp. 22-23 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 X1, X2, X3, and X4.
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
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 105 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
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
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 Na2HPO4, 1.47 mM KH2PO4, 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
(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.
3.
4.
5.
6.
7.
Aspirate media from wells and rinse each well with ~2 ml PBS.
Aspirate PBS. Add 1 ml of working concentration dye in growth media to
each well.
Incubate for 15 min at 37º C.
Prepare a glass slide with two 7 μl drops of Phenol-Red free media.
Aspirate dye containing media from wells and rinse each well with ~2 ml PBS.
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.
3.
4.
5.
6.
7.
Aspirate media from wells and rinse each well with ~2 ml PBS.
Aspirate PBS. Add 1 ml of working solution of dye in growth media to each well.
Incubate for 30 min at 37º C.
Prepare a glass slide with two 7 μl drops of Phenol-Red free media.
Aspirate dye containing media from wells and rinse each well with ~2 ml PBS.
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
(B1, D1 & F1): Vital staining with BODIPY-TR Ceramide
(Use a dipper and beakers to process coverslips)
You will be given BODIPY-TR Ceramide already at working concentration. The
essential aspect of this procedure is temperature. Follow the handling instructions
scrupulously to avoid high background.
1. Prepare a pre-cooled incubation chamber on ice by lining a square petri dish with
Parafilm. Pipette two 100μl drops of prepared 5μM ceramide-BSA onto the
Parafilm. Keep on ice while you are working with the chamber.
2. Remove coverslips from 6 well plate; place in a dipper in a beaker containing PBS
at room temp. Note which side of the coverslip has the cells bound to it. Remove
coverslips from the dipper, wicking off excess PBS and invert on the prepared
drops.
3. Incubate on ice for 30 min.
4. Prepare a beaker of PBS at 4oC. Place dipper in this beaker. Keep at 4oC until
needed.
5. Pipette 200 μl PBS at 4º C under the coverslip and lift the edge of the coverslip
slowly. (The extra buffer helps to prevent shearing of the cells off of the coverslip
due to surface tension.) Transfer coverslip to the dipper in 4o C PBS. Rinse by
gently dipping up and down 3X. This 4o C step is extremely important so don't
skip it.
6. Transfer the dipper to a beaker containing warmed growth media and incubate
for 30 min at 37º C. (Optional: Incubate cells in Hoechst dye at a concentration of 0.2
μg/ml by diluting a 1 mg/ml stock solution of Hoechst dye 1:5000 into the beaker of
growth medium)
7. Transfer the dipper to a beaker containing PBS at room temp.
8. Prepare a glass slide with two 7 μl drops of Phenol-Red free media.
9. Remove coverslips from the dipper, wicking off excess PBS. Mount on the
prepared glass slide. Seal with nail polish. View slides as soon as the nail polish
is dry.
(B2, D2 & F2): Vital staining with Rhodamine-Transferrin
(Use a dipper and beakers to process coverslips)
You will be given a 2 mg/ml stock solution of Rhodamine-transferrin. It is critical to
use the serum free media and the BSA containing media in the proper order for this
dye to work well.
1. Remove coverslips from 6 well plate; place in a dipper in a beaker containing PBS
at 37oC. Note which side of the coverslip has the cells.
2. Transfer the dipper to a beaker containing serum free media (DMEM) and
incubate for 20 min at 37º C.
3. Return dipper with coverslips to a beaker containing PBS.
4. Prepare a 100 μg/ml working solution of dye by diluting a 2 mg/ml Rhodaminetransferrin solution with DMEM + 1% BSA. Make enough for 100 μl per
coverslip.
____μl Stock solution + ___μl DMEM/BSA= ____μl working solution
19
5. Prepare an incubation chamber in a square petri dish lined with Parafilm. Pipette
two 100μl drops of working solution of Rhodamine-Transferrin.
6. Remove coverslips from the dipper, wicking off excess PBS and invert on the
prepared drops.
7. Incubate for 60 min at 37º C.
8. Pipette 200 μl PBS under each coverslip and lift. (This helps to prevent shearing of
the cells from the coverslip due to surface tension.) Transfer coverslips to the
dipper and place it in a beaker containing PBS.
9. Prepare a glass slide with two 7 μl drops of Phenol-Red free media.
10. Remove coverslips from the dipper, wicking off excess PBS. Mount on the
prepared glass slide. Seal with nail polish. View slides as soon as the nail polish
is dry.
Mounting Coverslips: The purpose of the nail polish is to prevent the sample from
drying out before you can look at it. The cells, of course, do not particularly like nail
polish either, but with a little care you can seal in many happy cells to capture their
images. Tack down the corners with a small gentle dab of nail polish. Once these are
hardened you can gently brush on polish around the edges. Once this has hardened
you should carefully rinse off the PBS salts with a wet Kimwipe. Your slide is now
ready for viewing. Remember your cells are alive and are best viewed as soon as
possible. Proceed through the sealing and viewing steps as quickly as you can
without exposing the microscope objective to wet nail polish!
Viewing slides and capturing images:
It is important that you view all slides (your own and others from the same table) on
the microscope. There should be 2 sets of results (plasmids U, X, Y and Z, each
counter-stained with 4 dyes). To save time, however, each person will only capture
images from his/her own slide. Find a representative (or “typical”) field. Without
moving the slide, you should capture an image using each fluorescent filter
appropriate for your sample (i.e., blue illumination/green emission for GFP, green
excitation/red emission for BODIPY-TR ceramide, MitoTracker Red, or rhodamine
transferrin, and UV excitation/blue emission for ER-Tracker Blue-White or Hoechst).
You might also want to capture a phase image of the same field of cells using
transmission light – this can help to provide more context for the fluorescent dyes.
Important: Colocalization studies using multicolor fluorescence staining techniques
are subject to potential artifacts. Since the images you are recording are all blackand-white, it is important to keep track of which image corresponds to which dye by
naming the images appropriately. A common complication is bleed-through of
strong fluorescent signals from one channel to another. Since fluorphores absorb and
emit light over a range of wavelengths, a very bright fluorophore can sometimes
fluoresce enough to be detected with the “wrong” filter set – e.g., you may see
fluorescence from the Hoechst dye in the green emission channel. In addition, cells
expressing very high levels of the GFP fusion protein may show GFP fluorescence
not only in the fluorescein (green emission) channel but also in the rhodamine
channel. Bleed-through fluorescence will usually look dimmer in the “wrong”
channel than the “right channel”, and you can also usually tell that it is bleedthrough because the color will look different to your eyes than authentic rhodamine
20
fluorescence (for example, GFP will usually look sort of yellowish-orange when
viewed through the rhodamine filter set, while rhodamine will look orange-red).
However, the camera doesn’t distinguish between different colors when it records
the image. Therefore, it is important for you to consider the possible contributions of
this type of artifact as you record your images and overlay them.
You will be determining the location of your fluorescent fusion proteins by
comparing the signals observed from the transgenic GFP expression with the signals
from the different vital stains. When you overlay these images, the GFP and vital
stain images should correspond very closely if and only if the dyes are localized to
the same organelle. Many cell structures are similar in general appearance. If your
slide gets slightly bumped between taking the different images, or you capture poor
quality or out-of-focus images, it may be very difficult to make unambiguous
decisions about which fluorophores colocalize in your cells. You should make an
effort to get a good and complete set of images, keeping in mind that you and the
other students in your section are working with living cells that will not look good
for long!
Save your images to a folder on the desktop. To minimize waiting time for the
microscopes, you should move your images to different computer for processing
and analysis. Colorize your individual images with red (rhodamine), green (GFP), or
blue (UV) colors (remember to indicate in your notebook how you colorized each of
the images). Then superimpose the images from a single field. Where the intensity
of red and blue images correspond, you will see magenta; red overlaid on green
looks yellow, and green plus blue looks turquoise when viewed on a computer
screen.
21