BioSci 145A lecture 11 page 1 © copyright Bruce Blumberg 2000. All rights reserved BioSci 145A...

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

• 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

How to get DNA into cells - introduction (contd)

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

– 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)

– 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

• 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

• 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)

How to make ES cells

• 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.

• 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

Targeting vector

electroporate

recombination

positive selection with dominant selective marker

negative selection to identify homologous recombinants

• 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

• 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

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