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How do I assess the quality of a cDNA library?
I want to clone CDS copies of genes from a library, but I don't know what's a typical expectation of getting a full length clone even for a shorter gene (~1000 bp). I know it doesn't have to be composed of full length cDNA for sequencing, but I'd imagine quality of starting materials is important there too.
Are there some common transcripts I can try to obtain from PCR that will tell me how fragmented the sequences are in there?
What's a typical result from a commercial cDNA library - are there any QC tests that are quick to run in the lab?
Getting Started with RNA-Sequencing (RNA-Seq)
Standard methods for RNA library preparation do not retain information on the DNA strand from which the RNA strand was transcribed. The ability to obtain information on the originating strand is useful for many reasons including the identification of antisense transcripts, determination of the transcribed strand of noncoding RNAs, and determination of expression levels of coding or non-coding overlapping transcripts. Overall, the ability to determine the originating strand can substantially enhance the value of a RNA-seq experiment.
Even beyond the decision to perform direction or non-directional library prep, several methods can be used to generate an RNA-seq library, and the details of these methods are dependent on the platform used for high-throughput sequencing. However, there are common steps.
- Abundant Transcript Removal: The majority of RNA molecules present in a cell are ribosomal RNA (rRNA), and since these are generally not of interest, they should be removed before making a library from the RNA of interest. Similarly, globin RNA is generally removed from blood samples and chloroplast RNA is often removed from plant leaf samples. Two popular options for this step are:
- Fragmentation: Fragments of an appropriate size for sequencing are generated by fragmentation of RNA prior to reverse transcription and cDNA synthesis, rather than by fragmentation of cDNA.
- Reverse transcription and second-strand cDNA synthesis: Complementary DNA (cDNA) is generated from the RNA template by a reverse transcriptase. This first strand cDNA is then made double stranded using a DNA polymerase.
- End repair, dA-Tailing and Adaptor Ligation: End repair of the ds cDNA library and optional dA-tailing (depending on the sequencing platform to be used) is followed by ligation to adaptors. The library is then ready for amplification and sequencing.
Important factors to consider when performing RNA-seq
Library preparation is an important part of the RNA-seq workflow and methods are currently available for library preparation for RNA-seq which offer simplified protocols and improved yields. However, the quality and accurate quantitation of input RNA still remains critical to ensuring successful cDNA synthesis and libraries. The following are some important factors to consider:
Identification and isolation of cDNA clones is important for the interpretation and functional investigation of sequenced genomes. cDNA sequence information facilitates the identification of exons and thus, the definition of the coding potential of genes. cDNA clones provide probes for a variety of studies of genomic expression, and clones that contain full open reading frames (ORFs) allow functional investigation of their protein products. The Mammalian Gene Collection project (MGC) was set up as a public domain program with the goal of providing full-length cDNA sequence information and physical clones for all genes, starting with the human and mouse genomes ( 1 ). The Riken Institute ( 2 ) and a number of private sector groups also have major programs aimed at collecting full-length clones. Initially, cDNA libraries in the MGC were prepared from a variety of cell lines and tissues. Clones randomly selected from these libraries were end-sequenced and unique clones thought to contain full ORFs were fully sequenced and made available for public distribution. The MGC has been highly successful. As of October 2004, >12 000 human and 10 000 mouse unique, full-length clones were reported to be sequenced and distributed, and an additional 3000 clones from each species evaluated as likely to contain full ORFs, are in the sequencing pipeline ( 3 ) ( http://mgc.nci.nih.gov/ ). However, as initially predicted, the yield of novel clones has decreased as the project has moved forward. Genes not represented by selected cDNA clones may have low expression levels and thus, be rare transcripts. Several approaches to isolation of rare clones are being used. One is generation and random sequencing of subtracted cDNA libraries. Another is prediction of mRNA sequences based on genomic sequence, and amplification of these specific sequences from cDNA using the PCR. A third approach is based on the idea, that transcripts that are rare in cDNA libraries prepared from available cell lines and easily obtained tissues, may be present at much greater relative abundance in specific tissues that can only be obtained in small amounts. In the brain, for example, there are well known instances of genes that are highly expressed in a relatively small numbers of neurons in highly discrete regions. For example, there are about 10 000 vasopressin containing cells in the rat paraventricular nucleus (divided between two major cell types) and 7000 in suprachiasmatic nucleus ( 4 , 5 ). A micro dissection that would ultimately yield a very small amount of RNA is required to obtain a sample enriched in these cells. Thus, methods of cDNA library production that are effective with very small amounts of starting material are important to obtain libraries with significant representation of these types of rare transcripts.
Most procedures for generation of cDNA libraries employ an oligonucleotide containing a short poly(dT) sequence as the primer in a reverse transcription reaction that uses purified poly-adenylated RNA as the template. Following a second-strand replacement reaction, synthetic oligonucleotide linkers or adaptors are ligated to the double-stranded cDNAs. These products, in many cases following restriction enzyme digestion, are ligated to an appropriately prepared vector. The adaptor/linker ligation is a relatively inefficient bi-molecular blunt-end ligation. It is performed with an excess of adaptor/linker, which has to be removed from the cDNA poduct before it can be introduced into the vector. This purification step results in loss of material. The subsequent ligation of the cDNA into vector, even in the case when ‘sticky ends’ are created, is a bimolecular reaction with limited efficiency. Incorporating the oligo-dT priming sequence into the vector to produce a ‘vector primer’, eliminates one or both of the ligation reactions and the purification step. Okayama ( 6 ) described a highly efficient vector-primer based cDNA library construction procedure in 1982. This method was not widely adopted, partly because it is technically demanding. Simpler vector-primer methods have been described previously ( 7 , 8 ), but they are also not widely used. Perhaps this is due to the fact that some technical limitations were not adequately addressed, or that commercial kits using oligonucleotide priming methods became widely available and easy to use. We have returned to a vector-primer method to generate cDNA libraries from very small amounts of tissue. We now present the evaluation of several cDNA libraries prepared from microgram quantities of total RNA.
Tips for two-hybrid screening
Minimizing false positives:
- Run replicates of the experiment to reduce the likelihood of indiscriminate reporter gene activation. Including a prey-only control also provides a baseline level of reporter activation.
- Vary the expression levels of bait and prey proteins - overexpression can lead to false interactions. Lowered expression levels will increase the stringency of your binding.
- And most importantly: Independently validate identified binding partners with other techniques!
Minimizing false negatives:
Interactions leading to detectable reporter signals depend on a number of factors, including protein expression and correct folding, post-translational modification, protein degradation, access to the nucleus in eukaryotic screens and fusion configuration. The possibility for problems with one or more of these parameters can lead to a high number of false negatives in Y2H screens.
- Testing your reporter system with a pair of proteins that are known to bind serves as a good positive control to ensure that your setup works. The last thing you want is to run your lovely library screen only to find the system itself is not working because your reporter protein doesn’t function in yeast!
- Many false negatives stem from or are exacerbated by expression of target proteins in heterologous systems (e.g. mammalian proteins in S. cerevisiae).
- While this may be difficult to resolve for a wide range of prey proteins, there are now a number of two-hybrid systems available in model organisms, including bacteria, alternative fungi ( C. albicans , pC2HB ) (7) and mammalian cells.
- If the problem is that your proteins are not post-translationally modified, co-expression of the enzyme responsible for the modification in the assay host strain can help.
- In order to reduce the chance that the configuration of your fusion proteins physically blocks the binding sites for the protein partner or the UAS/reporter gene, the same bait and prey libraries can be screened using both N- and C-terminal fusions of these proteins. This way both ‘ends’ of your protein are screened for binding.
- Using a variety of expression levels and systems can alleviate false negatives that arise from low or faulty expression of your target proteins. In fact, a good overall strategy to reduce false negatives and produce a more vigourous screen is to use a number of different bait and prey vectors. This has been shown to be as effective as using five independent protein interaction detection methods (6).
- In order to bind DNA and activate a reporter gene in eukaryotic cells, fusion proteins must gain access to the nucleus. To circumvent this requirement for transmembrane proteins, a split ubiquitin system has been devised (8).
- Comprehensive workflow for detecting coronavirus using Illumina benchtop systems application note.
- Sample preparation workflow:
- Viral RNA is extracted with the QIAGEN QIAmp Viral Mini Kit (Catalog No. 52904).
- Libraries are prepared with TruSeq Stranded Total RNA Library Prep Gold kit (Illumina, 48-sample kit, catalog no. 20020598 96-sample kit, Catalog No. 20020599).
- Samples prepared directly from swabs or similar sample sources are most suited for the NextSeq™ series of systems due to the recommended 10 million reads per sample.
- Libraries prepared from viral culture using the same library preparation workflow are well suited for other benchtop instruments including the iSeq™ 100, MiniSeq™, and MiSeq™ systems due to the lower recommended read count of 500,000 reads per sample.
- Local data analysis is performed with the Illumina Local Run Manager (LRM) Resequencing Module, with the SARS-CoV-2 reference genome.
- Downstream analyses can be performed with the IDbyDNA Explify platform (www.idbydna.com/explify-platform).
Enrichment-based sequencing with the Illumina DNA Prep with Enrichment (formerly known as Nextera™ Flex for Enrichment) library preparation workflow
- Enrichment workflow for detecting coronavirus using Illumina NGS systems application note
- Sample preparation workflow:
- Viral RNA is extracted with the QIAGEN QIAmp Viral Mini Kit (Catalog No. 52904).
- Viral RNA is reverse transcribed using Thermo Fisher's Scientific Maxima H Minus Double-Stranded cDNA Synthesis Kit (Thermo Scientific, Catalog No. K2561). Other reverse transcription workflows may be compatible, including the first strand synthesis module and second strand (nonstranded) synthesis module from NEB.
- Viral cDNA is used as input for the Illumina DNA Prep with Enrichment library preparation kit (formerly known as Nextera Flex for Enrichment), and enriched with the Respiratory Virus Oligo Panel (Illumina, Catalog No. 20042472).
- Sequencing is performed on the benchtop iSeq 100, MiniSeq, or MiSeq systems that are well suited for the low read requirements for these samples
- Local data analysis is performed with the Illumina Local Run Manager (LRM) Resequencing Module, with the SARS-CoV-2 reference genome.
- Downstream analyses can be performed with the IDbyDNA Explify platform (www.idbydna.com/explify-platform).
Amplicon sequencing with the AmpliSeq™ for Illumina SARS-CoV-2 Research Panel
- AmpliSeq for Illumina SARS-CoV-2 Research Panel product page
- Panel overview:
- This AmpliSeq community panel (made-to-order) contains 247 amplicons in two pools targeting the SARS-CoV-2 genome. The panel is designed for >99% coverage of the SARS-CoV-2 genome (
- Viral RNA is extracted with the QIAGEN QIAmp Viral Mini Kit (Catalog No. 52904).
- Refer to the AmpliSeq for Illumina Community Panel Reference Guide, Chapter 3, for the Protocol for RNA Panels.
- After library preparation, sequencing is typically performed on the benchtop sequencers, iSeq 100, MiSeq, MiniSeq, or NextSeq 500/550 due to the low read requirements for these samples
- The DNA Amplicon app in BaseSpace™ or the DRAGEN™ Metagenomics app can be used for data analysis.
This panel is intended for research use only (RUO) applications, and not for use in diagnostic procedures.
What's a good simple test for cDNA library quality? - Biology
Center for Molecular Medicine and Genetics
Wayne State University School of Medicine
540 E. Canfield
Detroit, MI 48201
EXAMINING THE FUNCTION OF PROTEINS AND PROTEIN NETWORKS WITH THE YEAST TWO-HYBRID SYSTEM
The yeast two-hybrid system provides a relatively straight forward approach to understanding protein function. Section II outlines the basic components of the interaction trap, a yeast two-hybrid system developed in the Brent lab (Gyuris et al., 1993). More detailed background information can be obtained in a number of recent reviews (Ausubel et al., 1987-1996 Finley and Brent, 1995 Mendelsohn and Brent, 1994). Section III contains an interactor hunt protocol, which is a condensed and updated version of the original protocol we first posted on the Internet in 1992 and subsequently updated (Finley and Brent, 1995 Finley et al., 1997). The version presented here is the one we currently use in our lab and represents our attempts to streamline and scale up the these techniques to facilitate characterization of large networks of interacting proteins. It is also useful for individual hunts. Section IV discusses alternative approaches specifically designed to look at large protein networks the ultimate goal of developing these and related approaches is to eventually map all of the interactions encoded by a genome. Section V discusses briefly two-hybrid approaches to understanding the functions of individual protein interactions.
Several different two-hybrid systems have been developed to study protein function. The garden-variety application is to learn about the function of a given protein by isolating proteins that interact with it, usually by screening a cDNA library. To conduct such an interactor hunt, a protein is expressed in yeast as a fusion to the DNA-binding domain of a transcription factor lacking a transcription activation domain. The DNA-binding fusion protein is generally called the bait . The yeast strain also contains one or more reporter genes with binding sites for the DNA-binding domain. To identify proteins that interact with the bait, a plasmid library that expresses cDNA-encoded proteins fused to a transcription activation domain is introduced into the strain. Interaction of a cDNA-encoded protein with the bait results in activation of the reporter genes, allowing cells containing the interactors to be identified.
The two-hybrid system developed in the Brent lab (the interaction trap) uses the E.coli protein LexA as the DNA-binding domain and a protein encoded by random E. coli sequences, the B42 "acid blob", as the transcription activation domain. Both proteins are expressed from multicopy (2µ) plasmids the LexA fusion, or bait, is expressed from a plasmid containing the HIS3 marker, and the activation domain fused protein, sometimes called the prey , is expressed from a plasmid containing the TRP1 marker. In the most commonly used bait plasmid, pEG202, the bait is expressed from the constitutive yeast ADH1 promoter. Related bait plasmids are available that express the bait fused to a nuclear localization signal. The most commonly used prey plasmid, pJG4-5, expresses proteins fused to the B42 activation domain, the SV40 nuclear localization signal, and an epitope tag derived from hemagglutinin, all driven by the yeast GAL1 promoter which is active only in yeast grown on galactose. Use of the GAL1 promoter to express the prey allows toxic proteins to be expressed transiently and helps eliminate many false positives in interactor hunts. The interaction trap uses two reporter genes that carry upstream LexA binding sites or operators: LEU2 and lacZ . The LEU2 reporters are integrated into the yeast genome the lacZ reporters typically reside on 2µ plasmids bearing the URA3 marker, though integrated versions are also available. Several versions of the LEU2 and lacZ reporters exists that have a range of sensitivities based on the number of upstream LexA operators. In general the LEU2 reporters are more sensitive to a given interacting pair of proteins than the lacZ reporters (Estojak et al., 1995) however, highly sensitive lacZ reporters have been used that contain several LexA operators and transcription terminator sequences downstream of the lacZ gene (S. Hanes, personal communication).
More details about the different strains and plasmids available for the interaction trap can be found elsewhere (Ausubel et al., 1987-1996 Brent et al., 1997 Estojak et al., 1995 Finley and Brent, 1994 Finley and Brent, 1995 Finley et al., 1997 Gyuris et al., 1993)
III. INTERACTOR HUNT PROTOCOL
Below I refer to typical strains and reporters needed for an interactor hunt. These include the sensitive LEU2 reporter strain EGY48, the sensitive lacZ reporter plasmid pSH18-34, a plasmid to express LexA fusions such as pEG202, and the library plasmid pJG4-5. The following is a condensed version of a previously published protocol (Finley and Brent, 1995). It is intended to clarify and expand on some important points in the original protocol. More details can be found at the web sites (Brent et al., 1997 Finley et al., 1997)
A. Testing baits Part 1: Does the bait activate transcription?
Before performing an interactor hunt it is very important to know the level of background activation by the bait protein itself. Almost every LexA fusion will activate the LEU2 reporter in EGY48 to some extent by itself. The amount of activation by a bait determines how, and whether, an interactor hunt is done. The most useful way to measure the level of activation is to determine the fraction of living cells that are able to grow in the absence of leucine (on leu- plates). Although it is not immediately obvious why a more strongly activating bait allows a larger fraction of EGY48 cells to grow in the absence of leucine, determination of this fraction is essential to performing an interactor hunt. The fraction can be represented as the number of colonies that grow on a leu- plate (Leu+ colonies) per living yeast cell plated. The number of living cells, or colony forming units (CFU), in an aliquot of cells is determined by plating dilutions on plates that contain leucine. Thus, the frequency of Leu+ colonies (or Leu+/CFU) is a ratio of the number of colonies that form on leu- plates over the number that form on plates that contain leucine. The test is done with the selection strain (the strain that already contains the lacZ reporter and bait plasmids) which is transformed with the empty library plasmid, pJG4-5 this closely mimics the conditions under which the selection for interactors will ultimately be performed. For a bait that is virtually unable to activate the LEU2 gene by itself, the frequency of Leu+ colonies in the test will be less than 10 -6 (i.e., less than 1 Leu+ colony will form when 10 6 CFU are plated on the leu- plates). Baits that activate a moderate level of transcription will result in Leu+ colonies at frequencies from 10 -4 to 10 -5 .
It is important to plate at least 10 6 CFU onto the leu- plates when testing a bait for activation of LEU2. To screen a typical library of 10 6 individual cDNAs, it will be necessary to plate over 10 6 CFU of the selection strain transformed with the library onto the leu- plates to select for interactors. If the background activation by a bait were tested by plating only 10 3 or 10 4 CFU onto leu- plates, and only one or a few Leu+ colonies form, it would be tempting to conclude that the bait activates LEU2 at a sufficiently low level to be used for an interactor hunt. However, if one were to then attempt to thoroughly screen a library of 3 x 10 6 individual cDNAs by plating over 3 x 10 6 CFU onto the leu- selection plates, at least 3000 colonies would form these would all be expected to be false positive (i.e., formed due to activation by the bait and not due to interaction of the bait with cDNA-encoded proteins). As discussed below, knowledge of the frequency of Leu+ colonies that arise from activation by the bait itself will also be important in determining the number of Leu+ colonies to pick for further analysis during an interactor hunt.
A second important test of the activating potential of a bait is its ability to activate the lacZ reporter. Generally, the most sensitive lacZ reporters (e.g., plasmid pSH18-34) are not as sensitive as the LEU2 reporters. In most cases a bait that produces Leu+ colonies at a frequency less than 10 -4 will not activate the lacZ gene, as measured by the ability of a colony to turn blue on an X-Gal plate. However, in rare instances and for unknown reasons, a bait that activates a very low level of the LEU2 reporter will activate the lacZ reporter to a significant level. Thus, it is essential to test for activation of the lacZ reporter when characterizing the bait.
Protocol 1 Testing whether a bait activates transcription
- Media recipes can be found at the web site (Finley et al., 1997) and elsewhere (Ausubel et al., 1987-1996 Finley and Brent, 1995 Guthrie and Fink, 1991).
- Liquid YPD media
- Liquid dropout media (Glu ura-, Glu ura-his-)
- Dropout plates (Glu ura-, Glu ura-his-, Glu ura-his- trp-, Gal/Raf ura-his- trp-, Gal/Raf ura-his- trp-leu-)
- X-Gal plates (Gal/Raf ura-his- trp- X-Gal)
- Yeast strain EGY48 (MAT µ ura3 his3 trp1 3LexAop- LEU2 :: leu2 ) or one of the less sensitive LEU2 reporter strains EGY191 or EGY189 (MAT µ ura3 his3 trp1 1LexAop- LEU2 :: leu2 ) (Estojak et al., 1995)
- The URA3 2 µ lacZ reporter plasmid pSH18-34, or a less sensitive lacZ reporter (Finley and Brent, 1995)
- HIS3 2 µ bait plasmid (e.g., a derivative of pEG202) expressing your bait protein fused to LexA
- Two control bait plasmids: one that encodes LexA fused to an activator like Gal4 as in the plasmid pSH17-4, and one that encodes a transcriptionally inert bait like LexA-Max (Zervos et al., 1993)
- The TRP1 2 µ library plasmid, pJG4-5, lacking cDNA
- See attached transformation protocol for additional reagents
1. Construct the selection strain either by serial transformation of EGY48 with pSH18-34 followed by your bait plasmid, or by co-transformation of EGY48 with your bait plasmid and pSH18-34. The selection strain (EGY48/pSH18-34/bait plasmid) should be grown on ura-his- medium in all subsequent steps to maintain selection for the bait and lacZ reporter plasmids. Pick three individual transformant colonies and streak to another Glu ura-his- plate for storage and later recovery. All three should behave identically in the tests below, in which case any one will serve as the selection strain into which the library will be introduced.
2. Transform the selection strain with pJG4-5 and select transformants on Glu ura-his-trp - plates. Take this opportunity to practice transforming the selection strain at high efficiency this will be necessary for transformation with the library DNA (Protocol 3).
3. Pick two or three transformant colonies and inoculate 10 ml liquid Glu ura-his-trp- medium (again, all colonies should behave the same, but performing the test on more than one can help ensure that the results are not due to some rogue mutant yeast or contaminant). Grow the liquid cultures at 30oC with shaking to OD600=1.0 (corresponding to about 107 cells/ml). This is mid-log phase, provided the culture started at OD600<0.2. If overnight cultures grow to a density greater than OD600=1.0, dilute to less than OD600=0.2 and then grow to OD600=1.0 so that the cells are in mid-log phase when harvested.
4. Make serial dilutions from 10-1 to 10-6 of each culture in sterile water.
5. Plate 100 m l of the culture and 100 m l each dilution onto two platesa:
6. Monitor the emergence of colonies during the next several days. Calculate the number of CFU that were plated on each Gal/Raf ura-his- trp-leu- plate by counting the number of colonies that form on the Gal/Raf ura-his-trp- plates. Calculate the number of Leu+ colonies/CFU. It is also worth taking note of the size of colonies after 2, 3, and 4 days (see below).
7. Test for lacZ expression. One way to do this is simply to patch individual transformants from step 2 to Gal/Raf ura-his- trp- X-Gal plates (about 1 cm x 1 cm patches) and incubate at 30oC. Yeast with a control LexA-activator fusion should turn blue overnight while those lacking LexA or containing a transcriptionally inert bait will remain white indefinitely. Alternatively, if the frequency of Leu+/CFU is higher than 10 -4 , it may be useful to replica plate from one of the Gal/Raf ura-his- trp-leu- plates (one with 200-500 colonies) to Gal/Raf ura-his- trp- X-Gal. This will reveal the frequency of blue colonies among the Leu+ colonies, a number that may be useful in determining hunt strategies (see below).
a Galactose is used in the medium because the actual selection will eventually be done on galactose plates to induce expression of the activation-tagged cDNA protein. Raffinose is added to aid yeast growth it provides a better carbon source than galactose alone but does not block the ability of galactose to induce the GAL1 promoter.
B. Testing baits Part 2: Does the bait protein enter the nucleus and bind to LexA operators in the reporters, and is the full-length fusion protein made?
There are rare reports of baits that are excluded from the yeast nucleus it usually possible to force these into the nucleus by including a nuclear localization domain N-terminal to LexA. Any small level of transcription activation by a bait could be taken as an indication that the bait protein enters the yeast nucleus. However, the ideal bait does not activate transcription, so another test is needed to show that it can occupy operators in the yeast nucleus. One simple test is the repression assay. This assay is based on the ability of most transcriptionally inert LexA fusions to inhibit transcription when bound to LexA operators situated between the TATA box and the upstream activating sequence (UAS) of a reporter. The reporter used for this test is the lacZ reporter in plasmid pJK101. This URA3 2 µ plasmid differs from pSH18-34 in that the GAL1 UAS is located upstream of the LexA operators. The GAL1 UAS activates the lacZ reporter at a high level in the presence of galactose, and for this particular derivative, it also activates at a low level in yeast grown on glucose. Any amount of repression of the GAL UAS by a bait, either in galactose or glucose, indicates that the bait enters the nucleus and occupies LexA operators.
Protocol 2 The repression assay
- Liquid YPD media
- Liquid dropout media (Glu ura-, Glu ura-his-)
- Dropout plates (Glu ura-, Glu ura-his-)
- X-Gal plates (Glu ura-his- X-Gal, Gal/Raf ura-his- X-Gal)
- Yeast strain EGY48 or a related strain
- The URA3 2 µ lacZ repression assay reporter plasmid pJK101
- HIS3 2 µ bait plasmid expressing your bait protein fused to LexA
- Two HIS3 2 µ control bait plasmids: one that encodes LexA fused to a transcriptionally inert protein, like Bicoid in pRFHM1, or LexA-Max (Zervos et al., 1993), and one that encodes no LexA, for example pRFHM0.
1. Transform EGY48 with pJK101 and select transformants on Glu ura- plates.
2. Combine three colonies from these plates and transform them with the HIS3 bait plasmid (and the HIS3 control plasmids). Select transformants on Glu ura-his- plates.
3. Pick four individual colonies from each transformation and streak a patch of them onto Glu ura-his- and Gal/Raf ura-his- plates containing X-Gal. Incubate at 30oC.
4. Examine the X-Gal plates after 1, 2, and 3 days. Yeast lacking LexA will begin to turn blue on the Gal/Raf plates after one day and will appear light blue on the glucose plates after two or more days. Yeast containing a bait that enters the nucleus and binds operators will turn blue more slowly than the yeast lacking LexA.
5. Baits that repress transcription of lacZ in pJK101 by 2-fold or less may not cause a visible reduction in blue on X-Gal plates. If no repression is observed on the X-Gal plates, perform the more sensitive liquid ß-galactosidase assays with transformants from step 2. Grow the transformants in 5 ml Glu ura-his- and Gal/Raf ura-his- liquid media, or on Glu ura-his- and Gal/Raf ura-his- plates for 2 days, before doing ß-galactosidase assays (Miller, 1972).
An ideal bait protein for an interactor hunt is one that does not itself activate transcription but does repress in the repression assay. It is also useful to verify that the full-length fusion protein is made. In some instances, proteases in yeast will cleave specific portions of a bait, leaving a truncated LexA fusion that still binds to operators. To demonstrate that the full-length bait protein is made one can perform a Western blot on extracts from yeast cells that harbor the bait plasmid, immunoblotting with either an antibody to LexA or one specific to the protein fused to LexA. The simplest way to do this is to prepare yeast cell extracts by growing yeast in liquid culture (lacking histidine to maintain selection for the bait plasmid) to OD 600 = 0.5, spinning 1 ml of the
culture to pellet the cells, and resuspending the cells in 50 m l of 2X Laemmli sample buffer (Laemmli, 1970). The cells can then be broken by freezing on dry ice followed by boiling for 5 min prior to loading on an SDS polyacrylamide gel (about 15 m l/lane). The proteins can then be transferred to a filter and blotted with standard immunoblotting (Western) methods (Ausubel et al., 1987-1996 Harlow and Lane, 1988).
C. Screening a library for interactors
Most cDNA libraries available for the Brent lab version of the yeast two-hybrid system contain over 10 6 individual cDNAs (in plasmid pJG4-5). In theory, a library with 10 6 individual cDNAs includes cDNAs for messages that were more frequent than 1 in 10 6 mRNA molecules in the mRNA population used to make the library. To have a chance at isolating the rarest cDNAs in a library, it is important to collect more yeast transformants than there are individual cDNAs in the library. Thus, for a library with 10 6 individual cDNAs, one might try to obtain 2-3 x 10 6 yeast transformants. With the most common yeast two-hybrid strains one can obtain up to 10 5 transformants per µg of library plasmid DNA using the attached transformation protocol.
A pilot transformation should be performed with the selection strain to determine the transformation efficiency that can be obtained. This allows one to calculate how many individual transformations to set up to obtain the desired number of total transformants. The transformation mixes are plated onto 22cm x 22cm Glu ura-his-trp- plates, attempting to get 1-2 x 10 5 transformants/plate. Again, the number of individual transformation mixes to put on each plate is calculated from the expected transformation efficiency derived from pilot experiments. The transformants are collected and stored frozen. Aliquots are then plated to ura-his-trp-leu- Gal/Raf plates to select interactors.
Protocol 3 Transforming the selection strain and selecting potential interactors
- Liquid dropout media (Glu ura-his-, Gal/Raf ura-his-trp-)
- Dropout plates (Glu ura-his-trp-, Gal/Raf ura-his-trp-leu-, Glu ura-his-trp-leu-)
- X-Gal plates (Glu ura-his-trp- X-Gal, Gal/Raf ura-his-trp- X-Gal)
- Sterile water
- Sterile glycerol solution (65% (v/v) glycerol, 0.1 M MgSO4, 25 mM Tris-HCl 7.4).
- Glass beads (4 mm diameter Fisher Scientific), sterilized by autoclaving.
- Sterile 50 ml Falcon tubes
- Sterile 50 ml round-bottom polypropylene centrifuge tubes
1. Using the selection strain prepared in Protocol 1, perform pilot transformations (as suggested in Protocol 1 step 2) to determine transformation efficiency.
2. Based on your transformation efficiency, calculate the number of transformations to obtain the desired number of total transformants (i.e., each transformation = 1 µg library DNA/50 µl of cells as described in the transformation protocol). Also, calculate the number of transformations to be plated on each 22cm x 22cm Glu ura-his-trp- plate to get 1-2 x 10 5 transformants/plate (e.g., if your efficiency in pilot experiments is 5 x 10 4 transformants/µg you should set up 2 transformations for each 22cm x 22cm plate).
3. Based on the above calculations, grow the appropriate amount of the selection strain in liquid Glu ura-his- medium and set up the necessary number of transformations (see attached transformation protocol).
4. After the heat shock, invert the tubes several times to mix - gently. Remove 10 µl from several of the transformation mixes and make three dilutions (10 -1 , 10 -2 and 10 -3 ) each in sterile water. Plate 100 m l of each dilution onto 100 mm diameter Glu ura-his-trp- plates and incubate at 30oC. This will allow the total number of transformants to be calculated.
5. Plate the remainder of the transformation mixes (less then 2 ml total/plate) onto 24cm X 24cm Glu ura-his-trp- plate. There is no need to spin the cells or remove the PEG. The medium in these plates should be at least 0.6 cm thick, level, and free of bubbles. To achieve an even distribution of cells, pour about 100 sterile glass beads (4 mm diameter) onto the plate with the cells. Gently roll the beads around the plate to distribute the transformation mix, then pour the beads off, or onto the next plate. This technique works best when the surface of the plates is not too wet so that the medium absorbs the transformation mix. To achieve this moisture content, put newly solidified plates into a laminar flow hood with the lids ajar for about 1 h before plating.
6. Incubate the plates at 30oC. Colonies should appear after about 24 h. Continue incubation until colonies are 1 - 2 mm in diameter, which should take a total of approximately 2 days.
7. Place the plates at 4oC for 2 - 4 hours to harden the agar. Using the long edge of a sterile 75mm x 50mm glass microscope slide (and sterile technique!), scrape the yeast from the plate. Try not to scrape any agar as this will interfere with pipetting. Collect the yeast from the glass slide by wiping it on the lip of a sterile 50 ml Falcon tube.
8. Wash the cells twice with 2 or 3 volumes of sterile TE. It may be necessary to split into two or more tubes to effectively pellet. It is best to pellet the cells each time in a sterile round bottom polypropylene tube at 2000 g for 4 min so they may be easily resuspended. The pellet volume for 500,000 transformants will be about 8 ml.
9. Resuspend the cells thoroughly by swirling in 1 pellet volume of sterile glycerol solution. Mix well by vortexing on low speed. Freeze 1 ml aliquots at -70oC.
10. Determine the plating efficiency by thawing an aliquot of library transformants and making serial dilutions in sterile water. Plate 100 m l of each dilution onto 100 mm diameter Gal/Raf ura-his-trp- plates. Count the colonies that grow after 2 - 3 days at 30oC. Represent the plating efficiency in c olony f orming u nits (CFU) per unit volume of frozen cells. Note: to save time one can estimate the plating efficiency as
108 CFU/100 m l, and immediately proceed to steps 11 and 12. Once the actual plating efficiency is known, calculate the number of CFU that were actually plated in steps 11 and 12.
11. Thaw a 1 ml aliquot of transformed yeast and dilute 10-fold into 9 ml Gal/Raf ura-his-trp- liquid medium. Incubate at 30oC with shaking for 6 to 8 h to induce the GAL1 promoter and expression of the library encoded proteins. Pellet the cells by centrifugation at 2000 g for 4 min at 20 - 25oC and resuspend in 10 ml sterile water.
13. Plate less than 106 CFU (determined from the plating efficiency test in step 10) onto each 100 mm diameter Gal/Raf ura-his-trp-leu- plates. To avoid overcrowding of Leu+ colonies, do not plate more CFU than are expected to produce
20 background Leu+/plate (as determined in Protocol 1). Incubate the selection plates at 30oC. Colonies should appear in 2 - 5 days. To keep the plates from drying out after two days, it may be helpful to put them in plastic bags or containers, or put parafilm around each plate.
14. Pick colonies (see discussion below for number to pick) with sterile toothpicks or applicator sticks and patch, or streak for single colonies, onto another Gal/Raf ura-his-trp-leu- plate. If the Leu+ colonies are closely spaced it will be necessary to streak purify to single colonies to separate the different Leu+ clones. Ideally the Leu+ yeast should be streaked for single colonies to isolate them from contaminating Leu- yeast. However, when there are large numbers of Leu+ colonies to pick, it may be inconvenient to streak purify every one in this case, growing patches on a second selection plate will at least enrich for the Leu+ cells.
15. To show that the Leu+ phenotype is galactose-dependent, patch (or replica plate) the Leu+ yeast onto Glu ura-his-trp- master plates to turn off the GAL1 promoter and stop expression of the activation-tagged cDNA protein. Grow at 30oC for about 24 h.
16. Replica the master plates to the following five plates, in order: 1. Gal/Raf ura-his-trp- X-Gal 2. Glu ura-his-trp- X-Gal 3. Glu ura-his-trp-leu- 4. Gal/Raf ura-his-trp-leu- 5. Glu ura-his-trp-. Incubate at 30oC and examine the results after 1, 2, and 3 days.
17. Pick only those yeast that are Leu+ on galactose but not glucose. Keep in mind that if Leu+ clones were not purified in step 14, some patches may be contaminated with background Leu+ yeast, which will not be galactose-dependent. The galactose-dependent Leu+ phenotype indicates that reporter activation depends on expression of the library protein. Further characterize these by isolating the library plasmid and determining the interaction specificity.
Alternate protocol - liquid selection and amplification of Trp+ library transformants. We have had some success at selecting and amplifying library transformants in liquid culture (M. Kolonin and R. Finley, unpublished). To do this, we dilute individual transformation mixes after heat shock (from Protocol 3 step 4) 50-fold into liquid Glu ura-his-trp- medium and grow shaking at 30 o C until the OD 600 is
2.0. The OD 600 of this culture begins at less than 0.2 and usually takes 30-48 hours to reach 2.0. We then harvest the cells and proceed as in Protocol 3 step 8. By removing aliquots immediately after dilution and before harvesting and plating on Gal/Raf-ura-his-trp- we have estimated that transformants are amplified approximately 100-fold in this procedure. This approach eliminates the cost and inconvenience of selecting transformants on plates. The disadvantage is that there is no reliable way to verify that library transformants are evenly amplified.
How many Leu+ colonies should be picked? When considering how many Leu+ colonies to pick at step 14 of Protocol 3, it is important to take into account the background frequency of Leu+ colonies that the bait itself produces (represented as Leu+ colonies/CFU), as determined in Protocol 1, and the total number of library transformants obtained. To completely screen all of the library transformants, the minimum number of Leu+ colonies one would need to pick and characterize can be estimated by:
# to pick > (# Leu+ colonies/CFU) X (total # of library transformants)
If, for example, the background for a given bait were 10 -5 Leu+ colonies/CFU, one would need to pick and characterize at least 10 colonies to screen through 10 6 library transformants. More to the point, the first 10 colonies picked would be expected to be background, so to get an interactor that is rare in the library one might need to pick and characterize 20 or 30 Leu+ colonies.
Should galactose-dependent Leu+ colonies that do not turn blue on the X-Gal plates be further characterized? Yes. Of the galactose-dependent positives, several different classes of Leu and lacZ phenotype are possible. For example:
Class I. galactose-dependent Leu+ galactose-dependent dark blue on X-Gal
Class II. galactose-dependent Leu+ galactose-dependent light blue on X-Gal
Class III. galactose-dependent Leu+ white on X-Gal
Many hunts will yield Leu+ colonies from each class. Often this indicates that at least three different interactors are represented among the positives. A common mistake is to concentrate on only the "strongest" class (Class I above) and ignore the "weaker" class (Class III) which can include biologically significant interactors (Finley et al., 1996).
The next step for the galactose-dependent positives is to isolate the library plasmid from each and re-introduce it into the selection strain to show that the putative interaction phenotype depends on the library plasmid and not on mutations in the yeast or reporter genes. This test can often be performed at the same time as the specificity test described below. If the library has been properly screened to exhaustion, each interactor cDNA should be represented more than once in the putative positives. cDNAs corresponding to abundant messages may have been isolated many times. To reduce the amount of work in subsequent steps it is useful to determine which yeast contain identical cDNAs. This can be easily done by performing PCR with primers flanking the cDNA insertion site using DNA template from a quick yeast miniprep (Finley and Brent, 1995). PCR products can be digested with HaeIII and AluI and run on an agarose gel to reveal unique restriction fragment patterns for each cDNA (Finley and Brent, 1995). One or two of each unique library plasmid can then be rescued in E.coli and used in the specificity test.
D. Determining the specificity of interactors
Many of the proteins identified in interactor hunts are non-specific interactors: they appear to interact with a number of different unrelated LexA fusions. Non-specific interactors are frequently isolated in hunts using unrelated baits. They can be identified and discarded by testing the ability of the cDNA-encoded proteins to interact with a handful of bait proteins unrelated to the original bait. cDNA-encoded proteins that interact only with the original bait and not with unrelated baits are considered true specific interactors. The specificity test can be performed by introducing rescued library plasmids into different selection strains that each harbor a different bait plasmid. Transformants are picked and patched onto a Glu ura-his-trp- plate and then replica plated to indicator plates as in Protocol 2 steps 15 and 16. This method of testing specificity can be somewhat cumbersome if a large number of different library plasmids were isolated, and if these are to be tested for interaction with several different baits. For this reason we use the interaction mating assay (Finley and Brent, 1994) to perform the specificity test, as described in Protocol 3.
Interestingly, the commonly isolated non-specific interactors, which include heat shock proteins, ribosomal proteins, proteasome subunits, and other proteins, are not isolated in every interactor hunt, and in fact do not appear to interact with every bait. This highlights the importance of using several different bait proteins to test the specificity of an interactor. For example, frequently a non-specific interactor will interact with just 30% of the bait proteins tested. If only one or a few bait proteins are tested, a non-specific interactor could appear to be specific.
Protocol 4 The interaction mating assay
- Rescued library plasmid DNA
- Liquid YPD medium
- Liquid dropout media (Glu ura-)
- YPD plates
- Dropout plates (Glu trp-, Glu ura-his-, Glu ura-his-trp-, Gal/Raf ura-his-trp-leu-, Glu ura-his-trp-leu-)
- X-Gal plates (Glu ura-his-trp- X-Gal, Gal/Raf ura-his-trp- X-Gal)
- Applicator sticks (e.g. FisherBrand 01-340), or toothpicks, sterilized by autoclaving.
- Replica plating apparatus and sterile velvets or filters.
- Yeast strain RFY231 (MAT a ura3his3 leu2 ::3LexAop- LEU2 trp1::hisG LYS2) or EGY48. Note: RFY231 is EGY48 with the trp1-1 allele deleted (R. Finley, unpublished).
- Bait strains: S. cerevisiae strain RFY206 (MATa ura3-52 his3Æ200 leu2-3 lys2Æ201 trp1::hisG ) transformed with a URA3 plasmid containing a lacZ reporter, such as pSH18-34, and various HIS3 bait plasmids, such as derivatives of pEG202 that produce different LexA fusions. Each bait strain will contain a different bait plasmid. One strain should contain the original bait used in the interactor hunt.
1. Transform yeast strain RFY231 with the rescued TRP1 library plasmids and select transformants on Glu trp- plates (if EGY48 is substituted for RFY231, more than one Trp+ transformant should be analyzed to ensure than a trp1-1 revertant has not been selected). As a control, transform RFY231 with a library plasmid pJG4-5 that has no cDNA insert.
2. Use sterile applicator sticks or toothpicks to streak individual RFY231 transformants onto standard 100 mm Glu trp- plates in parallel lines (see Figure 1). Streaks should be at least 3 mm wide and at least 5 mm apart, with the first streak starting about 15 mm from the edge of the plate. A 100 mm plate will hold up to 8 different bait strains. Include at least one streak of the transformants with the control plasmid (no cDNA). Create a duplicate plate of streaked RFY231 transformants for each plate of bait strains to be used.
3. Likewise, streak different bait strains in vertical parallel stripes on a Glu ura-his- plate. Create a duplicate plate of bait strains for each different plate of prey strains to be used. Incubate both sets of plates at 30oC until growth is heavy. When taken from reasonably fresh cultures (for example, plates that have been stored at 4oC for less than a month) streaked RFY206-derived bait strains take about 48 hours to grow and RFY231-derived strains take about 24 hours.
4. Print the RFY231 derivatives and the RFY206 derivatives onto the same replica filter or velvet so that the streaks from the two plates are perpendicular to each other (see Figure 1).
5. Lift the print of the two strains from the filter or velvet with a YPD plate. Incubate the YPD plate at 30oC overnight. Diploids will form where the two strains intersect. One strain may grow more rapidly than the other during this time but this does not hinder formation of diploids in the intersections.
6. Replica from the YPD plate to the following indicator plates, in order: 1. Gal/Raf ura-his-trp- X-Gal 2. Glu ura-his-trp- X-Gal 3. Glu ura-his-trp-leu- 4. Gal/Raf ura-his-trp-leu- 5. Glu ura-his-trp-. Incubate at 30oC and examine the results after 1, 2, and 3 days. Only diploids will grow on the X-Gal plates and only interactors will grow on galactose plates lacking leucine (Figure 1).
What next? Although the methods described above allow several types of false positive to be eliminated, they do not address the biological significance of the interactions observed. In some instances the sequence of a specific interactor will suggest that its interaction with the bait may have a real in vivo function. However, two-hybrid interactions can occur between proteins that normally do not interact (for example, because they are never expressed at the same time or in the same tissue or subcellular compartment). A good first step to show biological significance is to verify the interaction by a different, biochemical technique, preferably co-precipitation from a cell in which both proteins are expressed. Ideally, the next step would involve a functional assay for the new protein, to show, for example, that the new protein is involved in the same biological process as the bait protein. The following two sections include a few additional ways to address function.
IV. TWO-HYBRID METHODS TO STUDY LARGE SETS OF PROTEINS AND PROTEIN NETWORKS
Finding interacting partners can reveal much about the function of a protein. Most regulatory proteins, for example, appear to function by contacting other proteins. This is true for proteins that regulate many different cellular processes, including transcription, translation, DNA replication, signal transduction, cell cycling, differentiation, and programmed cell death. All of the proteins involved in a given process together can be thought of as a network of interacting proteins. The members of each interacting network are linked through protein-protein contacts. A complete understanding of any given process can only be achieved when all of the components of the protein network regulating it are identified. Yeast two-hybrid systems offer approaches to characterizing individual interactions and whole networks of proteins.
Isolating a new interacting protein can reveal information about function if the sequence of the new interactor indicates similarity or identity with a protein whose function has been at least partially characterized. However, it is still often the case that the sequence of a interacting protein reveals little about its function. Another approach is to assume that the new protein functions in the same network as the original bait protein and to use the new protein as a bait to identify other members of the network. Repeating this process increases the chances of isolating a previously characterized protein, or one whose sequence provides clues to function. In principle, this approach could be used repeatedly to isolate all of the components of a regulatory network. Because some regulatory proteins may be shared by different cellular processes (e.g. regulation of cell cycle and DNA replication by p21 CIP1 (Li et al., 1994)), and networks for many different processes may be connected (e.g. a signal transduction pathway and the activation of gene transcription), this approach could identify many expressed genes from a small number of starting points.
An approach complementary to performing sequential hunts is to use the interaction mating assay to look for interactions between increasingly large sets of proteins (Bartel et al., 1996 Finley and Brent, 1996). In one variation of this approach, large panels of baits are collected in baits strains placed on plates in grids (e.g., in the standard 96-well format). The grids can then be screened simultaneously for interactions with individual prey proteins. Bait strains can be created as described in Protocol 4 using bait plasmids that express various proteins of known and unknown function. Large panels of bait strains can be collected and stored frozen indefinitely and then screened against any number of prey strains.
One such collection contains over 700 different bait proteins from our own work and from numerous other labs that use the interaction trap. Screening a protein against such a panel enables one to quickly test its ability to interact with a large number of known proteins, most of which have been characterized to some extent, and have been chosen for study because of their known or suspected involvement in some biological process. Thus, finding an interaction between a tested protein and a member of the panel often gives an immediate clue about the biological function of both proteins. While the number of proteins in any such panel is far less than the number of proteins in a good library, this approach does offer the advantage of screening the test protein against a set of proteins enriched for those of current interest to the biological community. More restricted panels of bait proteins, for example those known or suspected to function in a particular pathway, or those isolated in sequential interactor hunts, can provide a useful resource for characterizing new proteins. Such a panel may also be useful to characterize differences in the patterns of interactions made by wild-type and mutant variants of proteins such as those created in vitro or associated with particular diseases or other phenotypes.
For some proteins, this approach offers additional advantages over screening a library using a traditional two-hybrid scheme. Proteins that activate transcription when fused to LexA or another DNA-binding domain can be difficult to use in conventional interactor hunts. Though methods are available to reduce the sensitivity of the reporter genes (Durfee et al., 1993 Estojak et al., 1995) it is not always possible to reduce the reporter sensitivity below the threshold of activation for some baits. Moreover, reduction in reporter sensitivity carries with it the risk that the reporters will not detect weakly interacting proteins. Thus, an alternative for proteins that activate transcription as baits, is to use them as preys to screen existing panels of baits, or even libraries of baits. Interaction mating approaches also have clear advantages for proteins that are somewhat toxic to yeast the prey vector allows conditional expression of toxic proteins in the presence of a bait, and often the interaction can be observed because the reporters are activated even if the cells subsequently become inviable.
V. TESTING THE FUNCTION OF INDIVIDUAL INTERACTIONS
Finding the position of a protein within a network of interacting proteins can provide information about the function of the protein and the network. However, ultimately, the nature of each individual protein-protein contact must be understood. Several two-hybrid methods allow the significance of individual protein interactions to be analyzed.
A. Mapping interaction domains
Determining the domains within a protein that are responsible for its interaction with other proteins can provide a valuable insight into the way a protein functions. Several approaches are available to map interaction domains with yeast two-hybrid methods. All start with a bait protein and prey protein that interact and activate the reporter genes. Derivatives of one of these proteins are constructed and tested for interaction with the other. We usually make derivatives of the prey protein because derivatives of the bait protein may differ in their ability to activate the reporters by themselves, which complicates interpretation of the results. Derivatives of the prey protein can be made and tested for interaction with the bait in several ways. In any approach it is important to keep in mind that the prey is a fusion to an N-terminal activation domain and must be maintained in the correct reading frame. One approach is to subclone restriction fragments encoding parts of the prey fusion protein into the prey vector (i.e., pJG4-5 or derivative) and introduce the resulting vectors individually into selection strains harboring the bait vector or control vectors. Alternatively, derivatives can be tested for interaction using the mating assay as described in Protocol 4. A second approach is to make N-terminal or C-terminal deletion derivatives of the prey fusion protein and test them for interaction with the bait, again by individual transformation into selection strains or by the mating assay. Deletion derivatives can be constructed in a cloning vector (Ausubel et al., 1987-1996), and then subcloned into the prey vector, pJG4-5. Alternatively, the deletion derivatives can be constructed directly in a derivative of the prey vector. For example, pZP4-5o and pJF3 are derivatives of pJG4-5 that have unique, rare restriction sites downstream of the cDNA cloning sites which allow C-terminal deletions to be constructed by unidirectional exonuclease III digestion from the 3' end of the insert (R. Finley, Z. Paroush, and J. Fonfara, unpublished). Similarly, pJF2 contains unique 5' restriction sites that allow N-terminal deletions to be constructed. A third approach is to make random DNA fragments encoding parts of the prey protein, for example by sonication (e.g., ref. (Stagljar et al., 1996), and insert these into the prey vector. Finally, a variety of techniques are available to make single and multiple point mutations of one interactor, which can then be inserted into the prey vector to test for interaction with a bait.
B. Construction of dominant negative mutants
A powerful approach to understanding protein function is to create and express dominant negative forms of the protein that inactivate the function of the wild-type version (Herskowitz, 1987). The yeast two-hybrid system provides a method to design and assay potential dominant negatives. One type of dominant negative is a protein mutated so that it still interacts with one of its protein partners but lacks other functional domains. In this case the "partner" could be another protein or the same protein if it forms homodimers. Expression of the mutant form of the protein might be expected to bind to the partner protein and make it inaccessible to the wild-type version. One way to create such a mutant is to isolate the minimal domain of a protein that will interact with another protein partner as described in the previous section. If the interacting domain is just a fraction of the protein it would be expected to lack other functional domains, and would therefore be a candidate dominant negative. A related but more precise approach could be used for proteins that have at least two different known partners. For example, if protein A interacts with both proteins B and C, mutant varieties of protein A could be constructed and tested in the two-hybrid assay for their ability to interact with just protein B but not protein C. In this case, we would have precise knowledge of the function missing in the dominant negative (interaction with protein C).
It is worth noting that, while the dominant negative effect is frequently open to multiple interpretations (Herskowitz, 1987), functional inferences from the type of dominant negatives referred to here may be less uncertain. This is because we know that the dominant negative interferes with a specific protein interaction we have designed it that way and tested it in the two-hybrid system.
C. Disrupting protein interactions
The yeast two-hybrid system provides an assay to develop reagents that disrupt protein interactions. Such reagents can be used in vivo to probe the function of individual protein interactions. Frequently a protein makes functional contacts with several other proteins. For example, the catalytic subunit of a protein kinase may interact with one or more regulatory subunits and with substrates. Deletion of the gene encoding the kinase could provide information about the function of the protein as a whole, but would not provide information about the individual interactions that it makes with other proteins. As mentioned in the previous section, certain types of dominant negative mutants may be created that interfere with specific interactions made by a wild-type protein. In the kinase example, a dominant negative kinase might be created that interacts with its regulatory subunit but not its substrate such a mutant would be expected to compete with the wild-type kinase for regulatory subunits.
Another type of potential disrupter of protein interactions that can be identified with the two-hybrid system is a peptide that interacts tightly and specifically with one of a pair of interacting proteins. Such peptides have been isolated from a random peptide library using the interaction trap yeast two-hybrid system as described by Colas et al. (Colas et al., 1996). These authors created a peptide library using a plasmid related to pJG4-5 that expressed random peptides fused to an activation domain and an inert platform molecule, E.coli thioredoxin. To find peptides that interact specifically with a bait protein an interactor hunt is performed as described in Protocol 2. Some of the specific peptides, called aptamers , would be expected to interact with surfaces of the bait that are required for interactions with other proteins. These are potential disrupters of specific protein interactions.
A two-hybrid assay can also be used to show that a potential disrupter can interfere with a protein-protein interaction. The two proteins can be expressed, one as a bait and one as a prey, and then the potential disrupter can be expressed to see if it reduces the ability of the bait and prey to interact and activate a reporter. We developed a method to test whether an interacting domain or a peptide aptamer can disrupt specific interactions (M. Kolonin and R. Finley, unpublished). A potential disrupter is first isolated as an interactor. The library plasmid expressing the potential disrupter is isolated and used to transform RFY231, and these transformants are mated with a special bait-prey interaction strain as described in Protocol 4. In this case, however, the bait strain expresses the original bait as a prey (activation domain fusion) from plasmid pMK1, and a protein that interacts with it as a bait. Disruption of the interaction results in loss of LEU2 transcription and inability to grow on leu- plates.
The methods outlined here present an integrated approach to understanding the function of proteins, protein interactions, and networks of proteins. First, all of the potential partners of a protein thought to be involved in a particular biological process can be identified. Second, many additional members of the same regulatory network can be identified in successive interactor hunts. Third, interaction domains can be mapped. Fourth, mutants incapable of specific interactions can be identified, and in many cases these mutants can be expressed in vivo to provide functional information. Finally, reagents can readily be developed that disrupt specific protein interactions, and then can be used to probe the function of these interactions in vivo .
I thank Mikhail Kolonin, Jennifer Fonfara, and Catherine Nelson, for providing comments, and Mikhail Kolonin and members of the Finley lab for contributions to the protocols. I also thank the members of the Brent lab for their many contributions to the protocols. I especially thank Roger Brent who co-wrote previous versions of the interactor hunt protocols.
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Brent, R., and al., e. (1997). http://xanadu.mgh.harvard.edu/brentlabhomepage4.html. web site.
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Finley, R. L., Jr., and Brent, R. (1994). Interaction mating reveals binary and ternary connections between Drosophila cell cycle regulators. Proc Natl Acad Sci U S A 91 , 12980-12984.
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Production of Vaccines, Antibiotics, and Hormones
Traditional vaccination strategies use weakened or inactive forms of microorganisms to mount the initial immune response. Modern techniques use the genes of microorganisms cloned into vectors to mass produce the desired antigen. Doctors then introduce the antigen into the body to stimulate the primary immune response and trigger immune memory. The medical field has used genes cloned from the influenza virus to combat the constantly changing strains of this virus.
Antibiotics are a biotechnological product. Microorganisms, such as fungi, naturally produce them to attain an advantage over bacterial populations. Cultivating and manipulating fungal cells produces antibodies.
Scientists used recombinant DNA technology to produce large-scale quantities of human insulin in E. coli as early as 1978. Previously, it was only possible to treat diabetes with pig insulin, which caused allergic reactions in humans because of differences in the gene product. In addition, doctors use human growth hormone (HGH) to treat growth disorders in children. Researchers cloned the HGH gene from a cDNA library and inserted it into E. coli cells by cloning it into a bacterial vector.
In Summary: Medicinal Biotechnology
Transgenic organisms possess DNA from a different species, usually generated by molecular cloning techniques. Vaccines, antibiotics, and hormones are examples of products obtained by recombinant DNA technology. Transgenic plants are usually created to improve characteristics of crop plants.
Biotechnology in agriculture can enhance resistance to disease, pest, and environmental stress, and improve both crop yield and quality.
Although several recombinant proteins used in medicine are successfully produced in bacteria, some proteins require a eukaryotic animal host for proper processing. For this reason, the desired genes are cloned and expressed in animals, such as sheep, goats, chickens, and mice. Animals that have been modified to express recombinant DNA are called transgenic animals. Several human proteins are expressed in the milk of transgenic sheep and goats, and some are expressed in the eggs of chickens. Mice have been used extensively for expressing and studying the effects of recombinant genes and mutations.
Figure 1. Corn, a major agricultural crop used to create products for a variety of industries, is often modified through plant biotechnology. (credit: Keith Weller, USDA)
Manipulating the DNA of plants (i.e., creating GMOs) has helped to create desirable traits, such as disease resistance, herbicide and pesticide resistance, better nutritional value, and better shelf-life (Figure 1). Plants are the most important source of food for the human population. Farmers developed ways to select for plant varieties with desirable traits long before modern-day biotechnology practices were established.
Plants that have received recombinant DNA from other species are called transgenic plants. Because they are not natural, transgenic plants and other GMOs are closely monitored by government agencies to ensure that they are fit for human consumption and do not endanger other plant and animal life. Because foreign genes can spread to other species in the environment, extensive testing is required to ensure ecological stability. Staples like corn, potatoes, and tomatoes were the first crop plants to be genetically engineered.
Transformation of Plants Using Agrobacterium tumefaciens
Gene transfer occurs naturally between species in microbial populations. Many viruses that cause human diseases, such as cancer, act by incorporating their DNA into the human genome. In plants, tumors caused by the bacterium Agrobacterium tumefaciens occur by transfer of DNA from the bacterium to the plant. Although the tumors do not kill the plants, they make the plants stunted and more susceptible to harsh environmental conditions. Many plants, such as walnuts, grapes, nut trees, and beets, are affected by A. tumefaciens. The artificial introduction of DNA into plant cells is more challenging than in animal cells because of the thick plant cell wall.
Researchers used the natural transfer of DNA from Agrobacterium to a plant host to introduce DNA fragments of their choice into plant hosts. In nature, the disease-causing A. tumefaciens have a set of plasmids, called the Ti plasmids (tumor-inducing plasmids), that contain genes for the production of tumors in plants. DNA from the Ti plasmid integrates into the infected plant cell&rsquos genome. Researchers manipulate the Ti plasmids to remove the tumor-causing genes and insert the desired DNA fragment for transfer into the plant genome. The Ti plasmids carry antibiotic resistance genes to aid selection and can be propagated in E. coli cells as well.
Flavr Savr Tomato
The first GM crop to be introduced into the market was the Flavr Savr Tomato produced in 1994. Antisense RNA technology was used to slow down the process of softening and rotting caused by fungal infections, which led to increased shelf life of the GM tomatoes. Additional genetic modification improved the flavor of this tomato. The Flavr Savr tomato did not successfully stay in the market because of problems maintaining and shipping the crop. However, since that time numerous crop plants have been developed and approved for sale and consumption. Corn, soybeans, and cotton in particular have been widely adopted by U.S. farmers.
Simple, rapid and reliable methods to obtain high quality RNA and genomic DNA from Quercus ilex L. leaves suitable for molecular biology studies
Isolation of high-quality RNA and genomic DNA (gDNA) from many samples is a necessary step before accomplishing molecular biology studies. The particular composition of Quercus ilex leaves, specially hard and rich in cell wall material, polyphenolics and secondary metabolites, usually results in preparations contaminated with non-nucleic acid compounds. Although many methods have been developed, each case of study demands a protocol adapted to the specific plant sample and the pursued research objectives. We have evaluated several protocols to establish the methodology that best suited to our current genetic and molecular studies on Q. ilex. Our priority was to select the simplest methods reducing the plant starting material and the time employed, without compromising yield, quality and integrity of the isolated nucleic acids. Our results point to two protocols based on silica-membrane purification, as the most convenient for Q. ilex leaf tissue, and both procedures are greatly improved by adding insoluble polyvinyl polypyrrolidone during the isolation process. The protocols optimized here can be completed at the microfuge scale and allow a researcher to process 48 samples in 1 h, producing high quality preparations suitable for the routinely molecular biology applications with higher efficiency than other more labour and time-consuming protocols.
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What's a good simple test for cDNA library quality? - Biology
Illumina offers three approaches to help researchers with sequencing of SARS-CoV-2. Next-generation sequencing (NGS) provides an effective way to screen samples and characterize viruses without prior knowledge of the infectious agent. The Illumina SARS-CoV-2 workflows described below are not intended for diagnostic purposes (this bulletin does not address the COVIDSeq workflow). Application notes that provide detailed information and example results are included for each workflow, below.
Total RNA sequencing with the TruSeq™ Stranded Total RNA Gold library preparation workflow