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EXAMINING THE FUNCTION OF PROTEINS AND PROTEIN NETWORKS WITH THE YEAST TWO-HYBRID SYSTEM

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EXAMINING THE FUNCTION OF PROTEINS AND PROTEIN NETWORKS WITH THE YEAST TWO-HYBRID SYSTEM

Russ Finley, April 1997


I. SUMMARY

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. 


II. INTRODUCTION

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 106 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 106 CFU onto the leu- plates when testing a bait for activation of LEU2. To screen a typical library of 106 individual cDNAs, it will be necessary to plate over 106 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 103 or 104 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 106 individual cDNAs by plating over 3 x 106 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

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Reagents

  • 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 (MATµura3 his3 trp1 3LexAop-LEU2::leu2) or one of the less sensitive LEU2 reporter strains EGY191 or EGY189 (MATµura3 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

Method

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 ml of the culture and 100 ml each dilution onto two platesa:

  • Gal/Raf ura-his-trp-
  • Gal/Raf ura-his-trp-leu-

Incubate at 30oC.

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.

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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

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Reagents

  • 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.

Method

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 OD600 = 0.5, spinning 1 ml of the

culture to pellet the cells, and resuspending the cells in 50 ml 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 ml/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 106 individual cDNAs (in plasmid pJG4-5). In theory, a library with 106 individual cDNAs includes cDNAs for messages that were more frequent than 1 in 106 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 106 individual cDNAs, one might try to obtain 2-3 x 106 yeast transformants. With the most common yeast two-hybrid strains one can obtain up to 105 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 105 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

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Reagents

  • 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

Method

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 105 transformants/plate (e.g., if your efficiency in pilot experiments is 5 x 104 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 ml 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 ml 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 colony forming units (CFU) per unit volume of frozen cells. Note: to save time one can estimate the plating efficiency as ~108 CFU/100 ml, 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 30oC until the OD600 is ~2.0. The OD600 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 106 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

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Equipment and Reagents

  • 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 (MATa 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.

Method

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).

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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.

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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 p21CIP1 (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.

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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). As illustrated in Figure 2, 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 (Figure 2).