Download Gene targeting (contd)

BioSci 145A Lecture 11 - 2/13/2001
Transgenic technology and its implications
• Topics we will cover today
– how to get the DNA into cultured cells.
• chemical transfection of cultured cells
• cell fusion (won’t discuss)
• liposome-mediated transfer
• cationic dendrimers
• virus mediated
• electroporation
– how to get DNA into embryos
• biolistic gene transfer
• microinjection
• electrotransformation
• viral infection
– transgenic technology
• standard transgenesis
• gene targeting
– Dr. La Morte will discuss single cell microinjection techniques on 3/1
How to get DNA into cells - introduction
• General terminology
– transformation refers to the uptake of foreign DNA, e.g. plasmid transformation of bacteria.
– Transfection, strictly speaking, refers to the transfer of viral DNA
– when referring to animal cells, we tend to use the term transfection to distinguish the transfer of DNA from the
“transformation” of a cancer cell.
• Transfection efficiency varies greatly from one type of cell line to another using any method.
– Must usually test several methods to determine which one works best for your cells and hands.
• Stable vs. transient transfections is also relevant.
– Supercoiled plasmids are best for transient transfections, linear best for stable transfections
– stable transfectants usually have single integration site with multiple copies integrated
– transient transfectants may replicate extrachromosomally.
• Observation is that cells that take up any DNA take up all DNA
– e.g. if cells take up one type of plasmid from the surroundings, they will take up all types
– enables co-transfection, introduction of multiple plasmids/cell
– this is a fundamental and indispensable tool
How to get DNA into cells - introduction (contd)
• Many claims of superior transfection efficiency are made by companies who sell reagents for transfection
– Caveat emptor, one San Diego company uses a competitor’s product in house instead of the reagent they promote.
– one of the largest profit margin items in the industry
• unless you own stock in a company selling the reagents, make your own whenever possible
Chemical transfection - Ca3(PO4)2
• W. Szybalski (a very famous microbiologist) decided to set up a system whereby mammalian cells could be induced to take
up DNA, much like bacteria - first successful report in 1962.
– To maximize success he also developed the HAT selection method.
– By analogy to bacterial transformation, it was discovered that successful DNA transfer was dependent on the formation
of a co-precipitate of DNA with calcium phosphate
• after the method was well understood in 1973, it became widely used
• Graham and van der Eb (1973) Virology 52, 456-467 is the classic reference.
• Chen and Okayama (1987) Mol Cell Biol 7, 2745-52 (very high efficiency variant)
• General principle is to form a precipitate of DNA that can be taken up by endocytosis
– Mix DNA, in phosphate buffer with CaCl2 at precise pH and an insoluble CaPO4 precipitate forms
• precision in pH is critical, alterations of as little as 0.01 pH units affect efficiency
• leave on cells several hours to overnight
• wash ppt off and add fresh medium
– OR add DNA and buffer to cells at low (3%) CO2.
• Ppt forms automatically over time
Chemical transfection - Ca3(PO4)2 (contd.)
• advantages
– very simple
– very inexpensive
•
– extensive literature
– works for most cell types
disadvantages
– adherent cells only
– some touch and experience required to get good precipitates
– not particularly efficient in many cell types
– many cells do not like adherent precipitate
– difficult to automate or perform as a high throughput method
Chemical transfection - DEAE dextran
• diethylaminoethyl (DEAE) modified dextran is a positively charged polymer
– many other charged polymers have been used with varying degrees of success and reproducibility
• PEI - polyethyleneimine
• poly-L-lysine
• roll your own
• DNA adheres to the polymer and remains soluble
• by some unknown means, the complex interacts with the cells and is taken up by endocytosis
• advantages
– may work in cells that are refractory to other methods
– gentle, not very toxic to cells
– works for cells in suspension
• disadvantages
– doesn’t work well in many cell types
– doesn’t work well for stable transfectants
– unclear mechanism of action makes optimization troublesome
– moderately expensive
– low throughput
Lipofection - liposome mediated transfection
• produce unilamellar liposomes and allow DNA to interact with them. Liposomes can be produced by:
– sonication
– extrusion through a small pore membrane
– dilution into aqueous medium
• mix with cells and allow to interact
• for a long time it was assumed that liposomes mediate fusion with cell membranes. However endocytosis is now known to be
the mechanism
• various formulations
– cationic lipids only, e.g. DOTAP
– mixture of cationic and neutral lipids, e.g. lipofectin (DOTMA:DOPE)
– phospholipids
– cholesterol-related lipids
• all work to some degree
• advantages
– very simple to perform and optimize - anyone can do it.
– easy to automate, high throughput
– reliable and reproducible
– stable and transient assays work well
– works well with many cell types and in vivo
• adherent and nonadherent
Lipofection - liposome mediated transfection (contd)
• disadvantages
– many formulations require use of serum free, or serum reduced medium for good efficiency
• all types that use neutral lipids
– some formulations are unstable to oxygen
• DOTAP and other unsaturated lipids
– variable toxicity necessitates careful optimization for many types (e.g. Lipofectin)
– VERY expensive to buy (but almost free to make)
– for example
• BMB-Roche sells 2 mg of DOTAP transfection reagent for $285. This is enough for ~6 96-well plates ($48/plate)
– 1 gram = $142,500
• pure DOTAP costs ~$400/gram from Avanti Polar Lipids. Time and material to make liposomes in vials about
doubles this cost. About $0.20/96-well plate
– Manufacturers lie quite a bit about the performance of their reagents due to the profit margins
• many do not work well, others not at all
Cationic dendrimer mediated transfection
• polycationic polymers of various densities and patterns (e.g. Superfect)
• interact with DNA to form complexes
• these interact with cells and are taken up by endocytosis
• advantages
– may be more efficient than liposomes
– stable and easy to use
– low toxicity
– automation friendly, high throughput
– suspension or adherent cells
• disadvantages
– expensive
– not readily possible to synthesize
Electroporation - electricity driven transfection
• principle is that brief, strong electrical pulse creates transient pores in the cell membrane that allows exchange of molecules
• cells and DNA are placed into a cuvette between two plates.
– High DC voltage(500+ V) applied as a pulse
• square wave form appears to work better than exponential decay (best for bacteria)
• possible optimizations are voltage, pulse length, wave form.
– Some experimentation with RF (radio frequency) pulses suggests greater efficiency
• but apparatus is not readily available
• advantages
– very efficient when it works
– quite effective at making stable transfectants (e.g. ES cells)
• disadvantages
– only works well for cells in suspension
• devices for transfecting adherent cells do not work very well and are cumbersome to clean
– kills cells very effectively
– expensive equipment and cuvettes
– extensive optimization
– very sensitive to salt concentrations
Viral infection
• infection is absolutely the highest efficiency method possible
– 100% infection is routine
• DNA to be expressed is cloned into a virus that can infect your favorite cell type - two general types of virus utilized
– retroviruses (RNA viruses), e.g. RSV
• tend to integrate
• can be insertional mutagens!
• Relatively small sized insert
• narrow host range
– large DNA viruses (adenovirus, vaccinia)
• extrachromosomal replication
• tend to have broad host specificity
• tend to be lytic
• large inserts are possible
• many viral genes are not required for infective virions
– nonessential genes are removed, thus allowing the virus to accommodate foreign DNA.
– Most such viruses requires a packaging strain to get infective virus particles
• primarily for biosafety
• field is primarily driven by gene therapy applications
– most current information found in gene therapy literature
Viral infection (contd)
• advantages
– efficiency
– simplicity of infection
• disadvantages
– not really feasible to introduce multiple constructs per cell. Best for introducing a single cloned gene that is to be
expressed highly
– at least P2 containment required for most viruses
• lots of hoops to jump through with institutional review boards (IRB)
• viral transfer of regulatory genes, or oncogenes is inherently dangerous and should be carefully monitored
• not so many old virologists
– host range specificity may not be adequate
– many viruses are lytic
– need for packaging cell lines
How to get DNA into cells - summary
• common feature of nearly all transfection methods is to form dense DNA complexes of small, uniform size
– 75-100 nm seems best
• how the complex is made may not matter much, many variations are possible (thousands of papers)
– size uniformity of particles is strongly related to efficiency of transfection
• needs to be optimized for the type of cells and requirements of each experiment
• which method is the best one for me?
– What is working in the lab or surrounding labs?
• Troubleshooting is rate limiting step in science
– liposomes and cationic dendrimers generally the best
• fast
• reproducible
• broad applicability
– if cost is a concern, either make your own liposomes or use calcium phosphate
– electroporation and viral infection have important utility but restricted applicability
• electroporation is great for cells in suspension
• viral infection is great for a single gene
• single cell microinjection is now feasible (Dr. La Morte)
– throughput is low
– uniform delivery ensures reproducibility
How to get DNA into embryos (other than mouse)
• Why would we want to do this anyway?
– Determine function of identified genes
– develop animal models for various diseases
– confer desirable property
• Choice of method depends on model system, developmental stage and outcome desired
– early embryos
• if cells are large than direct microinjection is possible (e.g. Xenopus and zebrafish)
• otherwise use methods below
– later embryos, cells are too small for direct microinjection
• biolistic gene transfer
• electrotransformation
• viral infection
• liposome-mediated transfer
• transgenic techniques - germline transmission
– must be using an appropriate system
• mouse
• Xenopus
• Drosophila
• zebrafish
• C. elegans
– not yet in chicken, most amphibians
Embryo microinjection
• Simple, direct way to get DNA, RNA or proteins into embryos
– primary application is embryos with large cells (xenopus, zebrafish)
• needles used are ~ 1 m diameter
• Xenopus microinjection takes two basic forms
– oocyte injection
– embryo injection
• oocyte injection
– oocytes are immature eggs, do not divide
– these are dissected from ovaries and can be used for various experiments
– DNA must be injected into the nucleus (germinal vesicle)
• transcription is possible
– RNA must be injected into the cytoplasm
• translation is very robust, can continue for long periods of time (days)
Embryo microinjection (contd)
• oocyte injection (contd)
– applications
• in vivo expression screening
– microinject pools of mRNA generated from libraries and evaluate function
– various channels, receptors and transporters identified this way
• protein expression system
• electrophysiology
– advantages
• long term expression of injected materials
• cells do not divide
• transcription is possible
• apparatus is relatively inexpensive
• easy to collect and store oocytes
• unhurried injections
– disadvantages
• cells do not divide
• not a developing system, limited questions
• nuclear and cytoplasmic injections may be required
– e.g. reporter gene must be put in nucleus, mRNA into cytoplasm
Embryo microinjection (contd)
• Embryo microinjection
– typically performed from 1-32 cell stage, depending on effect desired
– embryos divide and develop
• microinjected materials are mosaically distributed
– no transcription of injected DNA before MBT
• zygotic transcription begins at the midblastula stage
• by then, microinjected DNA is very mosaic
– transgenic approaches
– RNA is well translated but less stable than in oocytes (24-36 hrs max)
– applications
• misexpression of mRNAs
• injection of mutant mRNAs
• gain of function
• loss-of-function
– mRNAs encoding dominant negative mutants
– neutralizing antibodies
– “antisense” RNA?
– Morpholino antisense oligonucleotides
• Can target injected materials to particular tissues by using fate maps and blastomere injections at 32 cell stage
Embryo microinjection (contd)
• Embryo microinjection (contd)
– advantages
• very early stages can be manipulated
• targeted injections possible
• possible to combine molecular biology with experimental embryology
– disadvantages
• no early transcription
• mosaic inheritance
• embryos are dividing
– limited time window for injections
Virus-mediated transfer
• Just as with cultured cells, viral vectors may be used to express transgenes in embryos
– identical viruses are used (retroviruses and adenovirus)
– similar host range issues
• use of retroviruses may require use of virus-free eggs (extremely expensive since most chickens carry one strain or
other of RSV)
• clone gene of interest into viral vector
– package into virions
– concentrate and determine titer (infections particles/volume)
– microinject into embryo
• applications
– primary application is with chick embryo
•
advantages
– relatively efficient
• disadvantages
– no expression in early embryos!
– may be impossible to express some genes
• e.g. DN-RAR
– retroviruses do not stay at site of injection
– survival issues
– non-specific effects
Biolistic gene transfer
• Somewhat bizarre method developed for very difficult problems (plant cells)
• very small particles are coated with DNA
– blasted into target tissue
• gunpowder
• compressed air
• advantages
– works in systems that are refractory to other methods
• e.g. plant cells
• regenerating limbs
– not very difficult
• disadvantages
– equipment requirement
– not particularly efficient
• only a few % of target cells survive and take up DNA
– tissues must survive partial vacuum
Electroporation
• Just like cultured cells, tissues and embryos can be transfected with DNA by electric pulse
• typical setup consists of a pair of microelectrodes (usually needles) in close proximity.
– Maneuver this into close proximity of target, add DNA and zap
• applications
– primary use is with chick embryos
– some use of RF transfection in other embryos but not widely practiced or accepted
• advantages
– can work in very early embryos
– can target small areas relatively well
• unlike virus-mediated transfection, the DNA only gets into cells near the electrode
• disadvantages
– equipment requirement
• electrodes must be custom made
– plenty of “touch” is required
– not so many applications yet
• chick embryo
– potential of contamination with bacteria and molds
Transgenic technology
• Transgenesis is either not possible or not feasible in all model organisms
– typical model organisms of interest are:
• C. elegans
• Drosophila
• zebrafish
• axolotl
• Xenopus
• chicken
• mouse
– transgenic techniques are well developed in
• C. elegans
• Drosophila
• mouse
– becoming reasonably doable for
• Xenopus
• zebrafish
– not readily possible
• chicken
• axolotl
• targeted gene disruption only works in a few organisms
– mouse
– C. elegans
“Standard” transgenesis - mouse
• standard transgenesis
– this involved microinjecting DNA into a fertilized egg (mouse) or embryo (Drosophila)
• some fraction of embryos undergo integration of DNA into genome
• some fraction of these transmit the transgene in the germline
“Standard” transgenesis (contd)
• Each mouse that harbors a transgene and transmits it in the germline is a “founder”
– founders must be evaluated before proceeding to large scale breeding and analysis
• keeping mice is EXPENSIVE ~$1.00/cage/day.
– Multiple females can be caged together
– but males must be kept individually
• downstream analysis is very time consuming, tedious and expensive
• what would we like to know about a founder line?
– How many copies of the transgene are present?
• Prepare DNA from tails, do Southern analysis and compare with DNA standards
• Transgene copy number varies from 1 to several hundred
• Level of transgene expression is usually proportional to the number of copies
– is the transgene expressed? Transgenes are not equally active at all integration sites.
• Northern or Western analysis
– Western is best but requires an antibody.
» produce an antibody to the protein
» engineer the transgene to express myc, flag or other common epitope
– Northern is more commonly performed
“Standard” transgenesis (contd)
• what would we like to know about a founder line? (contd)
– is transgene expression as predicted?
• If the transgene is under the control of a tissue-specific promoter (e.g. its own), is it expressed in the correct tissue at
the correct time in development?
– Tissue Northern blots
– in situ hybridization
• If the transgene is expressed from a ubiquitous promoter, is it expressed ubiquitously?
– tissue Northerns
– quantitative RNA blotting
– RT-PCR
– is the transgene transmitted faithfully?
• Multiple tandem copies of the same sequences could be problematic
• are expression levels similar in progeny of founders?
– Same is desirable
– could be more or less, or even absent
“Standard” transgenesis (contd)
• Applications
– Transgenesis is a gain of function method
• doesn’t speak to necessity of a gene, unless a mutation is being rescued
– rescue of a mutation
– promoter analysis
• identify temporal or spatial requirements for expression
• verify function of suspected enhancer elements
– create models for dominant forms of human diseases
– identify effects of misexpression
• particularly with genes showing temporally or spatially restricted expression, e.g. Hox genes
• advantages of transgenic technology
– analysis is performed in vivo
• best test for gene regulation
– much less difficult than targeted disruption
– relatively high efficiency compared with targeting
• disadvantages
– gain of function
– no ability to target integration site
– no control over copy number
– injected DNA must contain all regulatory elements
– can’t study transgenes with dominant lethal phenotypes
Gene targeting
• Targeted disruption of genes is very desirable, wave of the future
– great to understand function of newly identified genes from genome projects
• produce a mutation and evaluate the requirements for your gene of interest
– good to create mouse models for human diseases
• knockout the same gene disrupted in a human and may be able to understand disease better and develop efficacious
treatments
•
excellent recent review is Müller (1999) Mechanisms of Development 82, 3-21.
•
enabling technology is embryonic stem (ES) cells
– these can be cultured but retain the ability to colonize the germ line
– essential for transmission of engineered mutations
– derived from inner cell mass of blastula stage embryos
– grown on lethally irradiated “feeder” cells which help to mimic the in vivo condition
• essential for maintaining phenotype of cells
Gene targeting (contd)
• ES cells are very touchy in culture
– lose ability to colonize germ line with time
– easily infected by “mysterious microorganisms” that inhibit ability to colonize germ line
• ko labs maintain separate hoods and incubators for ES cell work
– overall, ES cells depend critically on the culture conditions to keep them in an uncommitted, undifferentiated state that
allows colonization of the germ line.
Gene targeting (contd)
• technique
– isolate genomic clones spanning the gene of interest from an ES cell library
– construct a restriction map of the locus with particular emphasis on mapping the exons
– create a targeting construct with large genomic regions flanking the region to be disrupted
– an essential exon(s) must be disrupted such that no functional protein is produced from the gene
• this should be carefully tested in cell culture before mice are made
– it is often useful to design the construct such that a reporter gene is fused to the coding region of the protein
• this enables gene expression to be readily monitored and often provides new information about the gene’s expression
– dominant selectable marker is inserted within replacement region
– negative selection marker is located outside the region targeted to be replaced
– DNA is introduced by electroporation and colonies resistant to positive selection are selected.
– Integration positive cells are subjected to negative selection to distinguish homologous recombinants
• homologous recombinants lose this marker
Gene targeting (contd)
Gene targeting (contd)
Gene targeting (contd)
• Technique (contd)
– homologous recombination is verified by Southern blotting
– factors affecting targeting frequency
• length of homologous regions, more is better.
– 0.5 kb is minimum length for shortest arm
• isogenic DNA (ie, from the ES cells) used for targeting construct. Polymorphisms appear to matter
• locus targeted. This may result from differences in chromatin structure and accessibility
• problems and pitfalls
– incomplete knockouts, ie, protein function is not lost
• but such weak alleles may be informative
– alteration of expression of adjacent genes
• region removed may contain regulatory elements
• may remove unintended genes (e.g. on opposite strand)
– interference from selection cassette
• strong promoters driving these may cause phenotypes
• Applications
– creating loss-of-function alleles
– introducing subtle mutations
– chromosome engineering
Gene targeting (contd)
• Applications (contd)
– marking gene with reporter, enabling whole mount detection of expression pattern (knock-in)
• advantages
– can generate a true loss-of-function alleles
– precise control over integration sites
– prescreening of ES cells for phenotypes possible
– can also “knock in” genes
• disadvantages
– not trivial to set up
– may not be possible to study dominant lethal phenotypes
– non-specific embryonic lethality is common
– difficulties related to selection cassette