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Practical sessions of Gene Regulation

Hence our module this semester is all about gene regulation, so all the practical sessions will be discussing a number of important technologies which are being used in investigating gene regulation.

Practical 1: Transcription


As we have mentioned in the lectures, gene expression control could be achieved at the transcriptional stage (transcriptional gene regulation) or post-transcriptional. In this practical we will discuss methods of studying the transcriptional gene regulation. To identify factors affecting transcription you have to have an accurate and sensitive technique to quantify transcription. A wide number of different proteins coordinate to carry out transcription of a given gene. The presence of some of these proteins is essential to warranty a successful and complete transcription. In other words without one of them, transcription will be impossible. However, other proteins have assistant roles; therefore elimination of one of these proteins may have a little or no effect on transcription. Therefore, the transcription assay which is used to investigate the role of these assistant proteins has to be very sensitive to be able to detect small variations in transcription. Detecting alteration in transcription after inactivating or over-expressing a certain protein will show whether this protein is involved in the transcription machinery or not. Here we will discuss a transcription assay called "promoter-reporter plasmids".

Promoter-reporter plasmids
assay is one of the widely and traditionally used assays to detect transcription. This method is based on the transfection of reporter genes. Thus we need to know what a reporter gene is. Reporter genes are genes easily identified and measured, and selectable markers. Reporter genes are often used as an
indication of whether a certain gene has been expressed in the cell or an organism. Two reporter genes are very common in gene regulation research area; beta-galactosidase (LacZ) gene and chloramphenicol acetyltransferase (CAT) gene.

The LacZ gene is a common reporter in bacteria, which encodes the protein beta-galactosidase. This enzyme causes bacteria expressing the gene to appear blue when grown on a
medium that contains the substrate analog X-gal. On the other hand, the CAT gene confers resistance to the antibiotic chloramphenicol. Therefore, bacteria contains this gene will be the only cells to grow in chloramphenicol containing agar.

Applications of the promoter-reporter plasmid assay
1- If you need to find out the promoter of a given gene. In
this case you need to design a plasmid contains the full sequence of this gene plus one of the reporter genes discussed above. This plasmid will be called wild type (WT) because it contains the original sequence of the gene. In the meantime you need to design a number of other plasmids; all contains the same reporter gene as in the wild type but each plasmid contains a different copy of the gene under study. These different copies of the gene carry different mutations in the regions which could be potential promoter or start site of the
gene. These plasmids will be called mutant 1- n. All plasmids will be then transformed into E.coli and grown on a suitable selected medium depends on which reporter gene has been used. The change in the transcription level, compared to WT will be detected using either coloremetric assay in the case of
the LacZ gene or by quantifying live bacterial colonies in the case of the CAT gene. The mutant which shows the lowest level of transcription, compared to the WT, should be considered as the promoter or the start site of the gene under study.

2- Another application is investigating a potential transcription factor to a given gene with known promoter. In this case you will need to design one plasmid as a WT as discussed in the above application. This plasmid should contain one of the reporter genes plus the full correct sequence of the gene. Another plasmid (contains the same reporter gene as in the WT) will be designed in parallel, contains a mutation disrupting the transcription factor binding site in the promoter of the gene. One more plasmid, which does not have to include a reporter gene, will be prepared. This plasmid will contain the full correct sequence of the transcription factor under study. All
plasmids will be transformed into E.coli, cultivated on the appropriate medium and fluctuations in transcription will be detected as mentioned above. Transcription levels will be compared between WT and the mutant. If transcription has been abolished in the mutant case, this means that the transcription factor under study is important in the expression of the given gene and disruption of its binding to promoter inhibited the expression.

An alternative way to achieve the same goal is to create the mutation in the transcription factor itself rather than the gene. Mutations should be located in regions disrupt its binding to the promoter of the gene.

Practical 2: DNA-Protein binding
As discussed in details in the lectures, transcriptional gene expression control is carried out primarily by two main ways; either by histone posttranslational modifications or by chromatin remodelling. Both mechanisms affect gene regulation, either activation or inhibition, by altering the chromatin status. Histone tails modifications, as acetylation, methylation and phosphorylation, need interactions between the enzymes carrying the chemical group to be added (acetyl, methyl or phosphate group) and the chromatin which is represented here by histones. In the same way, the ATP-dependent chromatin remodelling complexes need to directly
interact with chromatin to perform its job in sliding or excising nucleosomes. Thus, this area of research was in an urgent need to develop a technology which could identify DNA-protein interactions. The main technique developed for that purpose is Chromatin Immunoprecipitaion (ChIP). ChIP could also play a nice role in investigating transcription factors as these factors have to bind to DNA to perform their jobs. In the next paragraph we will discuss breifiely the principles of the technique, while the attached video shows the detailed steps.
ChIP analysis entails the preservation of natural protein-DNA interactions formed in intact cells using either mild isolation procedures or macromolecular cross-linking reagents such as formaldehyde followed by chromatin fragmentation using sonication. Selective immunoprecipitation then is conducted using an antibody to the protein or protein modification of interest. Finally, reversal of the formaldehyde-induced cross-links, coupled with DNA isolation, then permits quantitative measurement of the precipitated DNA fragments using the technique of polymerase chain reaction (PCR) or quantitative PCR (qPCR). The abundance of a particular DNA fragment is correlated directly with the concentration of the mmunoprecipitated factor or modification originally associated with DNA at that site. In this manner, the level of DNA modifications, histone modifications, transcription factors, coregulatory modules, RNA processing enzymes, and other
related associations can be assessed following virtually any cellular treatment or perturbation. As useful as this technique has been, however, it was the ability to evaluate this precipitated DNA in a fully unbiased manner across defined chromosomal regions and eventually across the entire genome using oligonucleotide or DNA fragment–spotted tiled microarrays (ChIP-chip analysis), as outlined in Fig. 1, that increased significantly the biologic impact that ChIP analysis has had on transcription research. Interestingly, the utility of this particular genome-wide approach was exceptionally short-lived because the technique of massively parallel sequencing (eg, deep sequencing or next-generation sequencing) has rapidly become the method of choice for unbiased ChIP-derived DNA evaluation (Fig. 1). Of course, both these techniques rely on the availability of accurately sequenced rodent and human genomes. It is safe to say that chromatin immunoprecipitation linked to deep sequencing methods (ChIP-seq analysis) and the subsequent bioinformatic
analyses of these data sets are fundamentally altering transcription research. Both single genes and entire genomes now can be queried easily for features that range from detection of extensive epigenetic modifications to the basal or
induced appearance of a full assortment of regulatory transcription factors. The only absolute requirement for the technique is the availability of a fully characterized antibody. These methods, together with additional genome-wide
analyses such as DNase I or micrococcal nuclease predigestion, coupled with deep sequencing and with the emerging technique of quantitating gene expression via deep sequencing (RNA-seq analysis), are lending new vigor to the field of
transcription research, permitting full annotation of important genomes at both the epigenomic and regulomic levels.


















Fig1:
A diagram shows the principle of the ChIP.
Note: Please watch the video carefully and many times to
pick up the details of each step.






Practical 3: Nucleosome positioning
Nucleosome is the building unit of chromatin. As mentioned in details in the lectures, nucleosome positioning together with histone tails modification forms the histone code. Histone code determines the expression status of the gene. Therefore detecting nucleosome positioning is very important in investigating gene regulation. Nucleosome positioning
can be assessed by using, either low resolution analysis
through enzymatic or chemical cleavage of the linker region, or high resolution experiments by following the 10 bp cutting pattern generated by DNaseI (figure 2B).

A low resolution analysis of nucleosome positioning on the MMTV promoter is shown in figure 2C. This study compares digestion of naked DNA to chromatin with micrococcal nuclease. This nuclease cuts DNA preferentially in the linker
region between nucleosomes within the chromatin fiber. In the chromatin lane, the absence of the bands marked with a star after the micrococcal nuclease digestion and the presence of defined bands instead of a smear demonstrate that nucleosomes are precisely positioned over this promoter.

To further analyze the nucleosome distribution, a high resolution analysis was performed on the same promoter. Figure 2D shows a genomic footprint obtained after DNaseI digestion.








Figure 2. Nucleosome
positioning on the DNA. A. Precisely or randomly positioned nucleosomes. The vertical lines figure a restriction enzyme cutting site that will serve as a reference to map the nucleosomes. The linker region between the nucleosomes
can be cut either by the micrococcal nuclease, or by a chemical as MPE. B. The accessibility to DNaseI of DNAwrapped around a nucleosome is restricted. Only the region opposite to the core of the nucleosome can be cut. Rotational positioning of the nucleosome generates a periodic 10 bp cutting pattern. C.
Nucleosome mapping over the mouse mammary tumor virus (MMTV) long terminal repeat (LTR). Nuclei were isolated from a mouse cell line (904.13) containing 200 copies of a construct containing the MMTV LTR. The nuclei or naked DNA were
digested with micrococcal nuclease, the DNA was purified and digested with PstI. The DNA fragments were analyzed by Southern blotting, using the indirect end-labeling technique. The radioactive probe is marked by an arrow. The band on top of the two lanes is the PstI/PstI fragment. The other bands correspond to the micrococcal nuclease cutting sites. D. Nucleosome B may show rotational positioning. Nuclei from cell line S4 (CV1 cells containing MMTV LTR or naked DNA were digested with DNaseI, DNAs were purified and the cutting sites mapped using the LMPCR technique.

Another example for performing high resolution analysis, by testing DNAse hypersensitivity, is studying Chromatin organization of the promoter of the TFF1 gene. DNaseI hypersensitive sites (HS) were mapped in three human breast cancer cell lines, MDA MB231 (MDA), MCF7 (MCF) and HE5. In breast, TFF1 expression is induced by estradiol, via its nuclear receptor. In hormone-dependent breast cancers cells (MCF7) TFF1 is expressed, while in hormone independent breast cancers cells (MDA MB231, HE5) it is extinguished. Cells were untreated or treated for 2 h with estradiol. Nuclei were isolated and digested with DNaseI. The DNaseI hypersensitive sites were mapped using the indirect end-labeling technique . The DNA was digested with XbaI and the probe is shown by an arrow. The band on top is the XbaI/XbaI fragment. The other bands are DNaseI hypersensitive sites.

Results: the chromatin structure of the promoter of trefoil factor 1 gene (TFF1, formerly pS2) was analyzed in three human mammary cell lines that express (MCF7) or not (MDA MB231 and HE5) the gene. In MCF7 cells, addition of hormone
causes the appearance of two inducible DNaseI hypersensitive sites, HS1 and HS4. In contrast, in MDA MB 231 and HE5 cells lines only one constitutive DNaseI hypersensitive site, HS2, was observed. The presence of HS1 and HS4 is correlated with transcription, while the presence of the HS2 constitutive
DNaseI hypersensitive site can be correlated with a silent state of the gene.


Figure 3. Chromatin organization of the promoter of the TFF1 gene.







Practical 4: Ligation-mediated PCR (LM-PCR)
The regulation of gene expression at the transcriptional level is mediated in part by specific interactions between proteins and DNA. The binding of such transcriptional regulatory proteins to specific DNA target sequences modifies the reactivity of the latter toward treatments that cleave the DNA by chemical (KMn04, Fe-EDTA, DEP, OP2Cu+, ...) or enzymatic (DNAse I, micrococcal nuclease, restriction enzymes, exonuclease III, ...) means. In higher eukaryotes the principal difficlty in analyzing these interactions stems from the complexity of the genome. For example, in mammalian cells method for visualizing the cleavage sites should be sensitive enough to read a sequence (about 300bp) of a single copy gene present in a genome of about 3 billion bases (the complexity of mammalian genomes).

Several methods exist for such an analysis:
1) Separation of genomic DNA on sequencing gel, transfer to a membrane followed by hybridization of a highly radioactive probe. The other methods involve separation on a sequencing
gel of labeled material generated subsequently to solution hybridization of specific primers:

2) Single extension from a single primer.

3) Linear amplification by repeated extensions from a single primer.

4) Exponential amplification mediated by the ligation of a linker (LM-PCR).

The sensitivity of this latter method is such that starting from less than 1ug of genomic DNA, an analysis of 1/10th of the reaction products allows a visualization of the region of
interest in 2 hours. Here we will describe the latter method (LM-PCR).

Description of the Method of Exponential Amplification
The method is based on the exponential amplification between a known site and a large number of unknown nearby cleavage sites generated in vitro or in vivo by chemical or enzymatic means. In order to carry out the exponential amplification, a
synthetic linker is added to each of the unknown cleavage sites.

There are three steps to the method:
- Elongation of a first primer and ligation of the linker
- Exponential amplification with a second primer
- Probing of the amplified region with a third primer.

The three gene-specific oligonucleotides are chosen in order of increasing Tm which permits an increase in specificity at each step. The procedures from step to step as well as the choice of oligos, their Tm, their size and their position in the sequence is discussed in the following sections.


1) First primer elongation and linker ligation

A) Generation of blunt ends by primer elongation
In order to systematically ligate a linker at each cleavage site, it is necessary to render these sites compatible with this linker. This is achieved by generating double-stranded blunt ends at each cleavage site by elongation of a first primer (referred to as oligo no1). It is expected that blunt ends are generated essentially at the desired sites. Oligo no1 should hybridize to a region located up or downstream 150-200 bases from the putative binding site. After hybridization, elongation of oligo no1 is catalyzed by Sequenase TM. This is the enzyme of choice because it is highly processive and lacks exonuclease activity. Polymerization by Sequenase TM is carried out at 40oC. The Tm (2) of oligo no1 should theoretically be as close as possible to this temperature in order to minimize nonspecific hybridization. Furthermore, oligo no1 should be a minimum of 16 bases in length to ensure that the corresponding sequence is
statistically present only in the genome. In practice we select an oligo no 1 of 19 bases in length.

B) Ligation of the linker
The linker seq is for example in this case:
5'-GAATTCAGATC-3'
3'-CTTAAGTCTAGAGGGCCCAGTGGCG-5'
The linker is asymmetric and nonphosphorylated to prevent self-ligation.

2) Exponential amplification

Upon linker addition to the population of blunt ended fragments, there are two available sites between which an exponential amplification becomes possible. Starting from the products of the ligation reaction, the amplification takes place as follows :
a. Hybridization and elongation from oligo no2 using a thermoresistant DNA polymerase
b. Denaturation and hybridization of both oligo no2 and of the 25-mer linker
c. Elongation on both strands using the thermoresistant DNA polymerase
a. to c. : n cycles = 2n fragments

The amplification is performed using a thermoresistant DNA polymerase. The high temperature that can be used for the polymerization reaction promoting? The specificity of hybridization of the oligos (oligo no2 and the 25-mer strand on
the linker). Oligo no2 allows to increase the specificity of the reaction due to its high Tm (higher than that of oligo no1) and hybridizes 3' relative to the site of the hybridization of oligo no1. In general the site of hybridization of oligo no2 should contains one or two G/C at the 3' end of the oligo.
For each new oligo no2, different hybridization temperatures can be tested in order to obtain the best yield during the exponential amplification. This precaution can in fact be essential to the elimination of background due to nonspecific hybridization of oligo no2.

3) Probing the amplified region
To probe the amplified region, it is first necessary to denature the amplified DNAs and then hybridize fragments to oligo no3, radiolabeled at the 5' end using polynucleotide kinase. Maintaining the order of increasing Tm improves the
specificity of the reaction at each step. The Tm of oligo no3 should be thus higher than that of oligo no2, and oligo no3 should hybridize a site 3' to the site of hybridization of oligo no2. Furthermore, it is necessary that oligo no3 and not oligo no2 is used as a primer. The sensitivity of the method can be
considerably increased by carrying out several cycles of denaturation, hybridization, and polymerization.

To analyze several sequences of the same or different genes, it is possible to "multiprime" the genomic DNA by simultaneously elongating several oligo no1's at different regions and then by amplifying with several corresponding oligo no2's. In this case it is important : 1) to avoid
elongation of oligos no1 on two strands of the same seqence, and 2) to separate each oligo no1 by at least 400 bp. Multipriming confers the advantage of being able to analyze several regions from a DNA sample treated under eqivalent
conditions.




Practical 5: Gene expression detecting methods
(It will be described in the section)

End of the course.

Good Luck.
D. Mohamed Kamal