control of gene expression in...

Next Step MCAT Content Review: Biology and Biochemistry

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CHAPTER 35

Control of Gene Expression in Prokaryotes

A. INTRODUCTION

Genomic DNA represents the set of genetic instructions which ultimately govern all cellular activities.

In both prokaryotes and eukaryotes, these instructions must be implemented via the synthesis of RNA

and proteins. Thus, the portion of a cell’s genetically-inherited instruction set that is carried out at any

point in time is a function of the genes that are actively expressed. The first step in expression of a

gene, the transcription of DNA to RNA, is common to both prokaryotic and eukaryotic cells. In this

chapter, we will explore the mechanisms by which prokaryotic cells accomplish the regulation of gene

expression at the transcriptional level.

In most prokaryotes, the logic of transcriptional regulation is focused on the conservation of

limited cellular resources, which are often insufficient to support the transcription (and later

translation) of all of a cell’s structural genes. As a consequence, only genes encoding proteins that

support ongoing cellular functions are expressed continuously. The transcription of the remainder of a

cell’s structural genome is regulated in response to the needs of the cell. If a protein is not immediately

needed, its transcription will be suppressed. If a protein or a set of functionally related proteins is

required by a cell, then a signaling system will initiate transcription of the pertinent structural gene or

genes.

B. PROKARYOTIC TRANSCRIPTION

While prokaryotic transcription technically encompasses the transcription of both bacteria and

archaea, on the MCAT, and in most of molecular biology, it is synonymous with bacterial

transcription. The principle enzyme responsible for the synthesis of RNA in bacteria is bacterial RNA

polymerase, which catalyzes the polymerization of ribonucleoside 5’-triphosphates (NTPs). As in the

synthesis of DNA by DNA polymerase, RNA polymerase catalyzes the growth of an RNA polymer

exclusively in the 5’ to 3’ direction. Unlike DNA polymerase, RNA polymerase does not require a

preformed primer in order to initiate synthesis, but is initiated at specific sites at the beginning of

genes. This de novo process of initiation is a particularly important step in the regulation of prokaryotic

transcription.

Chapter 35: Control of Gene Expression in Prokaryotes

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I. RNA Polymerase Structure

The intact bacterial RNA polymerase is a metalloenzyme composed of two zinc molecules and five

different types of subunits, shown in Figure 35.1: α, β, β’, ω, and σ. The core catalytic subunit of the

enzyme, which consists of two α subunits as well as one β, one β’, and one ω subunit, is fully capable of

catalyzing the polymerization of NTPs.

Figure 35.1: Subunit composition of bacterial RNA polymerase

However, the core polymerase subunit does not bind specifically to the DNA sequences that

direct incoming polymerase enzymes to their correct position relative to the start site of transcription.

The σ subunit, which is weakly bound to the core subunit and is sometimes referred to as a sigma

factor, serves this purpose and is needed for the correct identification of the transcriptional start site.

This recognition and binding by the σ subunit is required for the synthesis of a complete, functional

mRNA molecule. Many bacteria are capable of producing different sigma factors, each of which

recognize different promoter regions. For example, E. coli can produce seven different factors that

initiate transcription of a factor-specific subset of genes. Some sigma factors, distinguished from one

another by their molecular weight, bind to the promoters of genes that encode proteins or RNA

molecules that are required for essential “housekeeping” processes. Other sigma factors direct RNA

polymerase to the promoters of genes that encode more specialized functions, such as proteins

required for adaptation to environmental stresses or those needed for the metabolism of nitrogen. The

relative rates of synthesis of specific sigma factors, then, represent one means by which bacterial cells

modulate transcription of specific genes, dependent upon the nature of the gene and on environmental

signals.


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II. Initiation

The DNA sequences to which the σ subunit of RNA polymerase bind when initiating transcription are

part of the promoter. These sequences are located approximately 10 and 35 nucleotides upstream of

the transcription initiation site, which is defined as the +1 position. Appropriately, sequences found in

these promoter regions are called the -10 and -35 elements. While these sequences are not identical in

all promoters, they are similar enough to establish consensus sequences, the bases most frequently

found at each position in many prokaryotic organisms. The six-nucleotide consensus sequence

associated with the -10 element, 5’-TATAAT-3’, is known as the Pribnow box, and is similar to the

TATA box found in eukaryotes. The initial binding between the polymerase and a promoter forms the

pre-initiation complex (PIC). The PIC is a closed-promoter complex, referring to the fact that the

DNA contained in the complex is not yet unwound. Binding of the RNA polymerase holoenzyme to

the promoter region of DNA and the formation of the PIC is shown in Figure 35.2.

Figure 35.2: Promoter identification and assembly of the PIC

After formation of the PIC, RNA polymerase then unwinds 12-14 bases of DNA from about -

12 to +2. This forms an open-promoter complex in which single-stranded DNA is available as a

template for transcription. Transcription is initiated by the β subunit. After the addition of

approximately 10 nucleotides in the form of NTPs, σ is released; the core subunit then moves along

the template DNA, carrying out elongation of the growing RNA chain. The transcription of a basic

prokaryotic gene is dependent on the strength of its promoter. In the absence of other regulatory

elements, a promoter’s sequence-based affinity for RNA polymerases varies, leading to the production

of differing amounts of transcripts depending upon the gene transcribed. The extent of that variable

affinity is a function of the degree of similarity between the nucleotide sequence of the promoter and


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the consensus sequence. As sequence similarity increases, so does the binding affinity of promoter for

RNA polymerase.

III. Elongation

During elongation, the polymerase remains associated with the template strand as it continuously

synthesizes mRNA, unwinding the template DNA ahead of it and rewinding the DNA behind it.

Within the unwound portion of DNA, eight to nine base pairs of the growing RNA chain are bound to

the complementary template strand of DNA. Structural analysis indicates that during elongation, the β

and β’ subunits form a structure that maintains the association between RNA polymerase and the

DNA template, while also forming a channel through which the template strand passes and in which

the polymerase active site can be found.

IV. Termination

Synthesis continues until the polymerase encounters a termination signal, at which point transcription

stops, the newly synthesized RNA strand is released from the polymerase, and the enzyme dissociates

from its DNA template. In E. coli – the bacterial model of prokaryotic transcription which you are

most likely to encounter on Test Day – there are two alternative mechanisms of terminating

transcription. The most common termination signal consists of a symmetrically inverted repeat of a

GC-rich sequence followed by approximately seven A residues. Transcription of the GC-rich inverted

repeat results in the formation of a segment of RNA that can form a stable stem-loop structure by

complementary base pairing. This self-complementary structure, known as the hairpin terminator,

interacts with the transcription factor NusA, disrupting the association between RNA polymerase and

the DNA template at the β subunit. This causes the termination of transcription. Because hydrogen

bonding between A and U is weaker than hydrogen bonding between G and C, the presence of A

residues downstream of the inverted repeat sequence of G and C is thought to facilitate the

dissociation of the more tightly bound, self-complementary G and C base pairs from their template.

This process is termed Rho-independent termination.

Alternatively, transcription can be terminated in some genes by a specific termination protein

called Rho, which binds extended segments of nucleotides found in single-stranded RNA. Since

mRNA in bacteria become associated with ribosomes and are translated while still being transcribed,

such extended regions of single-stranded RNA are exposed to binding by Rho only at the end of an

mRNA. This is known as Rho-dependent termination. Not surprisingly, the Rho-dependent

terminator occurs downstream of the translation stop codon and is composed of a non-repeating,

cytosine-rich sequence known as a Rho utilization site (rut) and a downstream transcription stop point

(tsp). rut serves as the mRNA binding site for Rho. Activation of Rho, which occurs upon binding rut,


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allows Rho to hydrolyze ATP. This hydrolysis powers Rho’s translocation along the mRNA strand,

which continues until Rho makes contact with RNA polymerase, which has stalled at tsp. At this point,

Rho disrupts the mRNA-DNA-RNA polymerase transcriptional complex through a mechanism

involving the allosteric effects of Rho on RNA polymerase. A schematic depicting Rho-dependent and

Rho-independent termination is shown in Figure 35.3.

Figure 35.3: (A) In Rho-independent termination, the terminating hairpin forms from

self-complementary sequences of the nascent mRNA. Upon interaction with NusA, it

promotes release of the primary transcript from the RNA polymerase-DNA complex. (B)

In Rho-dependent termination, Rho first binds to the rut site. Once activated, it

translocates downstream until it reaches the RNA polymerase complex, stimulating

release of the transcript.

C. NEGATIVE CONTROL OF TRANSCRIPTION

Transcription can be regulated at the stages of both initiation and elongation, but most transcriptional

regulation in bacteria operates at the level of initiation. An MCAT favorite for this content area is

gene regulation and the expression of genes involved in the metabolism of lactose, a carbon and

energy source, by E. coli. To conserve the cellular resources required for enzyme synthesis, the enzyme

that catalyzes the cleavage of lactose to glucose and galactose, β-galactosidase, along with other

enzymes involved in lactose metabolism, are expressed only when lactose is available for use by the

bacteria. In other words, lactose induces the synthesis of enzymes involved in its own metabolism. In

addition to requiring β-galactosidase, lactose metabolism involves the products of two other enzymes:

lactose permease, a transmembrane symporter that imports β-galactosidase into the cell, and a

transacetylase, which is thought to inactivate thiogalactosides that are transported into the cell along

with lactose by the permease enzyme.

I. Lac Operon


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The genes encoding β-galactosidase, permease, and transacetylase – lacZ, lacY, and lacA, respectively –

are expressed as a single contiguous unit in the chromosome. This arrangement is called an operon.

Generally, an operon is associated with a single promoter and is transcribed as a large, polycistronic

mRNA molecule. Transcription of the operon is controlled by the sequence of regulatory genes known

as the operator, which is adjacent to the transcription start site and immediately upstream of the

structural gene for β-galactosidase. The lacI gene encodes a protein that regulates transcription by

binding to the operator. Such a gene product is referred to as a repressor, which blocks transcription

when bound to the operator region. The repressor acts by binding to the operator in such a way as to

overlap with the promoter region, thereby preventing the binding or movement of RNA polymerase

on or along the DNA molecule. The structure of the operon is shown in Figure 35.4.

Figure 35.4: Organization of the lac operon

The presence of lactose leads to induction of the operon because the lactose gives rise to allolactose, a

metabolite, which is produced from the occasional transglycosylation of lactose by β-galactosidase.

Allolactose serves its effector function by binding the lactose repressor protein, the gene product of lacI,

thereby preventing it from binding the operator sequence of the regulated DNA. When regulated in

this way, the lac operon is referred to as an inducible system with allolactose as the inducer of

transcription. Lac operon induction and repression are shown in Figure 35.5.

Figure 35.5: 1. RNA polymerase 2. lac repressor protein 3. promoter 4. operator 5.

lactose 6. lacZ 7. lacY 8. lacA (A) The operon is in its “off” state, because allolactose is

unavailable to bind the lac repressor protein and decrease the protein’s affinity for the

operator. In the absence of allolactose-repressor binding, the lac repressor binds the

operator and obstructs transcription of the operon’s structural genes. (B) The lac operon

is in its “on” state. Allolactose, produced only when intracellular lactose is available,

binds to and decreases the affinity of the lac repressor protein for the operator. RNA

polymerase is able to associate with the promoter region of the operon and transcribe the

structural genes of the operon.


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II. Effectors and Repressors

In inducible systems such as the lac operon, binding of a low-molecular-weight compound, known as

an effector, to a particular repressor protein changes the conformation of the repressor. This change

reduces its affinity for the operator and allows transcription to proceed. In the lac operon, allolactose is

an effector molecule. In general, when the levels of effector are reduced, the repressor protein can bind

to the operator region for which it is specific, re-establishing the “off” state. In many cases, an operator

is specific for its particular operon; however, there are many examples of different operons with similar

operator sequences that are controlled by the same regulatory protein. The proteins encoded by these

operons are usually involved in related cellular processes.

III. Trp Operon

Another well-known example of prokaryotic transcriptional control is the trp operon, which modulates

the bacterial biosynthesis of tryptophan from its precursor molecule in E. coli. Unlike the lac operon,

which is an inducible system, the trp operon is a repressible system. The primary difference between a

repressible and an inducible system is the result that occurs when the effector molecule binds to the

repressor. In an inducible system, binding of the effector molecule to the repressor causes a

conformational change in the repressor which greatly decreases the repressor protein’s affinity for the

operator. Such action causes transcription of the operon to increase. In a repressible system, binding

by an effector molecule to the repressor gives rise to a conformational change that greatly increases the

repressor protein’s affinity for the operator, thereby causing transcription of the operon to cease.

The trp operon contains five structural genes: trpE, trpD, trpC, trpB, and trpA. These encode the

enzyme tryptophan synthetase, as well as several other enzymes involved in the pathway by which

tryptophan is synthesized from its precursor molecule, chorismic acid. The repressor of the trp operon

is produced upstream of the promoter by the constitutive, low-level expression of the trpR gene. In the

absence of tryptophan, the repressor, a tetrameric protein, is inactive. When tryptophan is present, it

binds to the repressor tetramer, causing a change in the repressor’s conformation that allows for the

association of the repressor with the operator region – located wholly within the operon’s promoter –

where it interferes with transcription. Effector molecules that serve as the activating ligand of repressor

proteins are known as corepressors. Their binding to the inactive repressor causes it to undergo a

conformational change, enabling the repressor-corepressor complex to bind to its corresponding

region and inactivate transcription of the structural genes of the operon. In general, the repressor

alone cannot bind the operator; when the concentration of the corepressor decreases, the “on” state

for the operon resumes. In the case of the trp operon, tryptophan acts as a corepressor for its own


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biosynthesis, providing a mechanism for feedback regulation in the synthesis of tryptophan. The

structure and regulation of the trp operon is shown in Figure 35.6.

Figure 35.6: Structure of the trp operon

IV. Attenuation

Another means by which the trp operon is regulated is attenuation. While the mechanism of repressible

expression responds to changes in intracellular tryptophan concentration, attenuation is responsive to

changes in the concentration of charged tRNA. Rather than decreasing gene expression by altering

the initiation of transcription, attenuation affects the process of transcription after it has begun.

One element of the operon is the leader sequence contained within the trpL gene, just

upstream of the first structural gene of the operon, trpE. This sequence contains four domains,

numbered 1-4, which are each partially complementary to one another. Domain 3 of the mRNA

synthesized from the gene can base pair with either domain 2 or domain 4. If domains 3 and 4 pair, a

stem-and-loop structure forms, interrupting further transcription. As discussed previously, such a

transcription termination sequence is rich in guanine and cytosine, and is followed by several uracil

residues which form weaker hydrogen bonds with adenine residues. Once the structure is formed,

RNA polymerase dissociates from the template DNA strand and the structural genes of the operon are

not transcribed. This 3-4 pairing occurs when the level of intracellular tryptophan is high. When

domains 2 and 3 pair, the stem and loop structure does not form and the enzymes required for

tryptophan biosynthesis are produced. This takes place when tryptophan levels within the cell are low.

The leader sequence codes for two adjacent tryptophan residues. When tryptophan levels are

low, the ribosome stalls while awaiting the delivering of a rare tryptophan-charged tRNA molecule.

While it is stalled, the ribosome obstructs the 1 domain of the transcript, preventing the formation of a

1-2 secondary structure. Domain 2 then is free to hybridize with domain 3. Formation of this 2-3


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secondary structure prevents formation of the 3-4 termination structure, as domain 3 is no longer

available to pair with domain 4. This mechanism of transcriptional attenuation is shown in Figure

35.7.

Figure 35.7: Mechanism of transcriptional attention of the trp operon. (A) Formation of

the stem-loop termination structure occurs when the intracellular tryptophan

concentration is high. (B) Formation of an alternate stem-loop structure, which occurs

when the intracellular tryptophan concentration is low, permits the continuation of

transcription.

D. POSITIVE CONTROL OF TRANSCRIPTION

The central principle of gene regulation exemplified by the lactose operon is that control of

transcription is mediated by the interaction of regulatory subunits with DNA sequences. This general

mode of regulation is applicable to both prokaryotic and eukaryotic cells. Regulatory elements like the

operator are called cis-acting control elements because they affect the expression of only linked genes


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on the same DNA molecule. Conversely, proteins like the repressor are known as trans-acting factors

because they can affect the expression of genes located on other chromosomes within the cell. Binding

of the lactose repressor is an example of a trans-acting factor involved in negative control. This,

however, is not universally true; many trans-acting factors are activators, rather than inhibitors, of

transcription.

The best-studied mechanism of positive control in E. coli involves the effect of glucose on the

expression of genes that encode enzymes involved in the catabolism of sugars, including lactose, that

provide alternative sources of carbon or energy for the bacterial cell. When glucose is present, it is

utilized preferentially by E. coli, and as long as glucose is available, enzymes involved in the catabolism

of other sugars are not expressed. As an example, when E. coli are grown in a medium containing both

glucose and lactose, the lac operon is not induced and only glucose is metabolized by the cell. This is

because glucose acts to repress the lac operon, even in the presence of its normal inducer, lactose.

Repression by glucose, an example of what is called catabolite repression, is now known to be

mediated by a positive control system coupled to the levels of cyclic AMP (cAMP). In bacteria, the

enzyme adenylyl cyclase, which converts ATP to cAMP, is regulated in such a way that when glucose

levels decline, cAMP concentration increases. cAMP acts to bind a transcriptional regulatory protein

called catabolite activator protein (CAP). The binding of cAMP to CAP permits binding of the cAMP-

CAP complex to its target DNA sequences, which in the lac operon are approximately 60 base pairs

upstream of the transcription start site. CAP then interacts with the α subunit of RNA polymerase,

facilitating the binding of polymerase to the promoter and activating transcription.

I. Dual Regulation of Carbohydrate Metabolism in E. coli

The combined regulation of lactose metabolism, by both the negative inducible and positive inducible

systems, causes the enzymes of lactose metabolism to be made in small quantities when both glucose

and lactose are present. The presence of glucose suppresses expression of CAP; when this occurs,

expression of the lac operon is due solely to the binding and inhibition of the lactose repressor protein

by lactose. Thus, when both lactose and the preferred carbon source, glucose, are present, there is little

expression of the enzymes of lactose catabolism. What expression does occur is known as “leaky

expression”; this provides for a basal level of catabolic enzymes to help process lactose when cellular

glucose is expended, but before lac operon expression is fully activated. Figure 35.8 summarizes the

expression of the lac operon under different environmental conditions.


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Figure 35.8: Structure of the lac operon (above), and lac operon gene expression under

different environmental conditions (below). P is the promoter region; O is the operator.


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Chapter 35 Problems

Passage 35.1 (Questions 1-6)

Mutant strains of E. coli that are deficient in regulation of the genes involved in lactose metabolism

were studied. These mutants were of two types. Constitutive mutants expressed all genes normally

involved in lactose metabolism, even when lactose was unavailable, while noninducible mutants failed

to express these genes in the presence or absence of lactose. Genetic mapping localized these

regulatory mutants to two distinct loci, called o and i, with o located immediately upstream of the

structural gene for β-galactosidase, z. Mutations affecting o resulted in constitutive expression;

mutations of i were either constitutive or noninducible.

The function of these regulatory genes was investigated by experiments in which two strains of

bacteria were mated, resulting in diploid cells containing genes from both parents. The inducibility of

z upon addition of allolactose to the medium containing the diploid offspring was observed. The

matings, as well as the inducibility of z in the offspring, are shown in Figure 1.


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Figure 1: Regulation of β-galactosidase expression in diploid E. coli mutants. (Note: i+ and o+ are

normal; i- and oc are mutant alleles; z1 and z2 are structural gene mutations with normal function.)

From these experiments, the lac operon model was proposed in which i encodes a repressor protein

and o functions as an operator.

1. Which of the following conclusions is LEAST supported by the results of the experiments in which

mutations in o and i were combined with different mutations in the structural genes?

A. In an oc/o+ cell, only structural genes that are physically linked to both oc and o+ are constitutively

expressed.

B. In an oc/o+ cell, only structural genes that are physically linked to oc are constitutively expressed.

C. In an i+/i- cell, structural genes located on the same chromosome as i+ are normally expressed.

D. In an i+/i- cell, structural genes located on the same chromosome as i- are normally expressed.

2. In terms of the lac operon model, which of the following is true of the i gene identified in the

passage?

A. Allolactose is unable to bind to some of the repressor protein synthesized in i+/i- cells.

B. i+/i- cells synthesize sufficient functional repressor protein to display normal inducibility.

C. The i+ gene encodes a nonfunctional repressor protein.

D. The i allele encodes the operator region of the operon.


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3. Which of the following is most likely to describe the oc mutant in cells containing only a single,

normal z allele?

A. It is dominant to o+ and is a cis-acting control element.

B. It is dominant to o+ and is a trans-acting control element.

C. It is recessive to o+ and is a cis-acting control element.

D. It is recessive to o+ and is a trans-acting control element.

4. Molecular analysis has identified several changes in the nucleotide sequence of oc when compared

to the sequence of o+. This sequence change is most likely to affect the binding of oc by:

A. bacterial RNA polymerase.

B. a repressor.

C. DNA polymerase.

D. an inducer.

5. Experimenters conducting the studies described in the passage concluded that the mutant i allele is

recessive to the wild type. Which of the following explanations is most consistent with this conclusion?

A. The copy of the lac operon adjacent to the defective i gene is activated by the protein product of

the wild-type i gene.

B. The genotype of a cell carrying one mutant and one wild-type operator site permits synthesis of lac

structural genes.

C. Haploid cells containing the wild-type i gene are inducible.

D. The repressor is a small protein that can diffuse within the cell and inactivate expression of

structural genes linked to i-.

6. Researchers developed a method to select for regulatory mutants by mating haploid strains

carrying two complete wild-type copies of the operon and the operon’s regulatory elements. Each

strain contained one defect, a single copy of either i- or oc. Which of the following correctly describes

the diploid offspring?

A. Repressor mutants display normal inducibility.

B. Repressor mutants display the same phenotype as diploids homozygous for mutant operator

alleles.

C. A mutation in one copy of the operator gene results in constitutive expression of an operon.


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D. A mutation in one copy of the operator gene confers the same inducibility phenotype as a

mutation in one copy of the repressor gene.

The following questions are NOT based on a descriptive passage.

7. The cya gene in E. coli encodes the enzyme adenylate cyclase, which produces cAMP. In a cya

mutant where intracellular cAMP concentration is decreased, which of the following is correct?

I. Expression of lacY is reduced.

II. Less catabolite activator protein will be found as a complex bound to the lac promoter.

III. Activation of the lac operon will be enhanced.

A. I only

B. II only

C. I and II only

D. I, II and III

8. The repressor in the trp operon:

A. is part of an inducible feedback mechanism.

B. is produced downstream of the trpR gene.

C. is constitutively expressed at a low level.

D. is in its active state unless bound by tryptophan.

9. Rifampin is a bactericidal antibiotic used in the treatment of tuberculosis. Its mechanism of action

involves the binding and inactivation of the β subunit of bacterial DNA-dependent RNA polymerase.

Which of the following actions in bacterial transcription is LEAST likely to be affected by disrupting β

subunit function?

A. formation of the channel in the RNA polymerase holoenzyme through which the DNA template

strand passes

B. catalysis of NTP addition to the RNA transcript

C. Rho-independent termination

D. identification of the transcription start site


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10. Which statement correctly relates the regulatory responses of E. coli to changes in intracellular

tryptophan levels?

A. Attenuation regulates initiation of trp operon structural gene expression.

B. Formation of the stem-loop transcription termination signal occurs when tryptophan

concentration is low.

C. Attenuation is responsive to changes in the intracellular concentration of tRNATrp.

D. Association between domains 2 and 3 of the trp leader sequence forms the stem-loop termination

structure.


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Chapter 35 Solutions

1. A.

Figure 1 shows that of the two diploid offspring, structural genes in the oc/o+ cell are constitutively

expressed if and only if the gene is physically linked to oc. This is seen in the constitutive expression of

z2, which is located on the same chromosome as, and structurally linked to, only oc. The constitutive or

inducible expression of oc and oi parent bacteria, respectively, further support this observation. This

contradicts the statement in choice A, the correct answer, and is consistent with choice B. Choices C

and D are also true statements. In i+/i- diploids, structural genes physically linked to either i+ or i- are

inducible.

2. B.

The gene i is responsible for the synthesis of a repressor protein. According to the passage, mutations

affecting i gave rise to either constitutive or noninducible expression of the operon. In cases where a

mutation in i gives rise to a repressor protein that is unable to bind the operator, structural genes of the

operon will be constitutively expressed. Conversely, if a mutation in i gives rise to a repressor protein

that can bind the operator, but can’t be bound by its effector, allolactose, expression of structural genes

of the operon cannot be induced. The constitutive expression of structural genes in the i- mutant

shown in Figure 1 shows that the mutation must be one that prevents binding of the repressor protein

to the operator. Since structural genes in i+/i- diploid offspring demonstrate an inducible phenotype, a

single wild-type copy of the gene must provide sufficient functional repressor to normally bind the

operator sequence of both genes, giving rise to normal operon inducibility in the offspring. This is

choice B. Choice A is contradicted by the previous discussion—the mutation i- appears to affect

binding of the repressor to operator, not of allolactose to repressor. Choices C and D are also

incorrect. The i+ gene encodes a functional repressor protein.

3. A.

Figure 1 shows that expression of z2 does constitutively occur in oc/o+ diploids (albeit at a lower level

compared to wild-type homozygotes). This indicates that oc is dominant to o+. Furthermore, regulatory

elements like the operator are called cis-acting control elements, because they affect the expression of

only linked genes on the same DNA molecule. Choice A, then, is the correct choice. Conversely,

proteins like the repressor are known as trans-acting factors, because they can affect the expression of

genes located on other chromosomes within the cell.


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4. B.

Figure 1 shows that the expression of structural genes in the i- mutant is constitutive. Thus, the

mutation must be one that prevents binding of the repressor protein to the operator. This is choice B.

5. D.

The inducible phenotype of structural genes in i+/i- diploid offspring demonstrates that a single wild-

type copy of the gene provides sufficient functional repressor to diffuse within the cell and bind the

operator sequence of both genes. This gives rise to normal operon inducibility in the offspring and is

consistent with choice D. While choice A is nearly a correct statement, it incorrectly indicates that lac

operon expression is activated, rather than inactivated, by the protein product of the wild-type i gene.

Choice B, while a true statement, doesn’t draw a distinction between inducible and constitutive

expression of the lac operon, both of which can occur depending upon the genotype of the cell—

expression of structural genes will occurs in both cases. Choice C, while also a true statement, concerns

expression in haploid cells and doesn’t relate expression patterns when both wild type and i- are

present.

6. C.

The discussion of previous questions explained that in diploid mutants, all structural genes linked to a

wild-type operator sequence are expressed when a single wild-type copy of i+ is available. Strains

carrying two copies of the operon’s regulatory elements will express normal inducibility if their single

defect is one copy of i-. This eliminates choices A and B. Regardless of the presence of functional

repressor protein, carrying a copy of oc will result in the constitutive expression of the structural genes

linked to that operator sequence. This is choice C. Choice D is false; a mutation in one copy of the

operator gene will result in the inducible expression of one operon and the constitutive expression of

the other. If only a single copy of the repressor allele is mutated, both operons will show normal

inducibility.

7. C.

In bacteria, the enzyme adenylyl cyclase, encoded by cya, converts ATP to cAMP. This process is

regulated in such a way that when glucose levels decline, cAMP concentration increases. cAMP acts to

bind a transcriptional regulatory protein called catabolite activator protein (CAP). The binding of

cAMP to CAP permits binding of the cAMP-CAP complex to its target DNA sequences. CAP then

interacts with the α subunit of RNA polymerase, facilitating the binding of polymerase to the promoter

and activating transcription of the operon structural genes. In the absence of adenylate cyclase,

encoded by cya, cAMP levels will not rise; CAP will be unable to bind and promote transcription of lac

operon structural genes, including that of lacY. Roman numeral I is then true, as is Roman numeral II.


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However, activation of the lac operon will be reduced rather than enhanced as a consequence of these

changes. Roman numeral III is thus false. This is choice C.

8. C.

The repressor in the trp operon is constitutively expressed at a low level by a gene upstream of the

promoter. Choice C, then, is correct and choice B is incorrect. The trp operon is a classic example of a

repressible (i.e. default on) system (eliminating choice A). In a repressible system, binding by an

effector molecule to the repressor protein gives rise to a conformational change that greatly increases

the repressor’s affinity for the operator, thereby causing transcription of the operon to cease. For this

reason, the repressor is in its inactive state unless bound by its co-repressor, tryptophan. Choice D is

therefore incorrect.

9. D.

The core RNA polymerase subunit does not bind specifically to the DNA sequences that direct the

incoming polymerase enzyme to its correct position relative to the start site of transcription. The σ

subunit, which is weakly bound to the core enzyme, serves this purpose and is needed for the correct

identification of the transcriptional start site. This process is independent of the function of the core

enzyme, which includes the β subunit. Choice D is the correct answer. During elongation, the β and β’

subunits form a structure that maintains the association between RNA polymerase and the DNA

template, while also forming a channel through which the template DNA strand passes and in which

the polymerase active site can be found. This eliminates choices A and B. During Rho-independent

termination, interaction of the hairpin terminator with the transcription factor NusA is mediated by

the polymerase β subunit; this interaction disrupts the association between RNA polymerase and the

DNA template at the β subunit, causing the termination of transcription. Choice C can be eliminated.

10. C.

Attenuation is responsive to changes in the concentration of charged tRNA, specifically that of

tRNATrp. This differs from the action of the repressor protein in the mechanism of repressible

expression, which responds to changes in intracellular tryptophan concentration. Choice C is correct.

Attenuation does not alter the initiation of operon transcription; it affects the process of transcription

once it has begun. Choice A is incorrect. Formation of the stem-loop transcription termination signal

by the association of domains 3 and 4 inhibits transcription of the trp operon, and occurs when

tryptophan concentration is low and the enzymes required for tryptophan synthesis aren’t needed by

the cell. Choices B and D are incorrect.


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CHAPTER 36

Eukaryotic Chromosome Organization and Control of Gene

Expression

A. INTRODUCTION

The basic mechanism of transcription is common to prokaryotes and eukaryotes. However, the

specific processes in eukaryotes are considerably more complex than those in bacteria and other

prokaryotes. This can be seen in three important differences between the prokaryotic and eukaryotic

transcriptional machinery:

• In bacteria, all genes are transcribed by a single RNA polymerase; in eukaryotes, more than one

type of RNA polymerase is responsible for the transcription of genes.

• Eukaryotic RNA polymerases require additional proteins to initiate and regulate transcription.

• Transcription in prokaryotes occurs on free DNA. In eukaryotes, transcription occurs on

chromatin, and regulation of chromatin structure strongly influences the transcriptional activity

carried out on eukaryotic genes.

The increased sophistication of regulatory options available in the eukaryotic cell likely evolved to

facilitate the complexity of directing the activity of the many different cell types of multicellular

organisms.

B. EUKARYOTIC RNA POLYMERASES

Eukaryotic cells contain three distinct nuclear RNA polymerases that transcribe different types of

genes. RNA polymerase I transcribes only the three largest species of ribosomal RNA (rRNA), which

are designated 28S, 18S and 5.8S, according to their rates of sedimentation during centrifugation.

RNA polymerase II encodes protein-coding genes, producing messenger RNA (mRNA), as well as

rRNA, transfer RNA (tRNA) and microRNAs, which are regulators of both transcription and

translation in eukaryotic cells. Some of the small RNAs involved in splicing and protein transport,

Chapter 36: Eukaryotic Chromosome Organization and Control of Gene Expression

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abbreviated as snRNA and scRNA, respectively, are transcribed by RNA polymerase II, while others

are transcribed by RNA polymerase III. Additionally, RNA polymerase III transcribes genes encoding

tRNA, as well as the smallest species of rRNA, 5S rRNA. RNA polymerases, which structurally

resemble bacterial RNA polymerases, specifically transcribe the organellar DNA in chloroplasts and

mitochondria. The classes of genes transcribed by eukaryotic RNA polymerases are summarized in

Table 36.1.

RNA synthesized RNA polymerase

mRNA II

tRNA III

5.8S, 18S, 28S rRNA I

5S rRNA III

snRNA, scRNA II and III

Mitochondrial genes Mitochondrial

Chloroplast genes Chloroplast

Table 36.1: Genes transcribed by RNA polymerases

All three eukaryotic RNA polymerases contain nine highly conserved subunits, five of which are

related to the core subunits of bacterial RNA polymerase. The structural homology of the core

catalytic subunits of eukaryotic and bacterial RNA polymerases suggest that all RNA polymerases

utilize similarly conserved mechanisms of transcription.

C. DNA BINDING PROTEINS AND GENERAL TRANSCRIPTION FACTORS

Specific proteins, called transcription factors, are required for RNA polymerase II to initiate

transcription. General transcription factors are involved in transcription from all polymerase II

promoters. For this reason, they represent a basic component of eukaryotic transcription. Additional

gene-specific transcription factors, discussed later in this chapter, bind to DNA sequences that control

expression of individual genes and are thus responsible for the regulation of gene expression. The

promoters of many genes transcribed by polymerase II contain a sequence similar to TATAA located

25 to 30 nucleotides upstream of the transcription start site. This sequence, referred to as the TATA

box, resembles the -10 sequence of bacterial promoters. The promoters of many genes transcribed by

RNA polymerase II also contain a second sequence element, called the initiator (Inr) sequence, which


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spans the transcription start site. While many promoters bound by RNA polymerase II contain both of

these elements, some contain only a TATA box and others contain only an Inr element. A large

number of those promoters that lack a TATA box but contain an Inr element also contain an

additional downstream promoter element (DPE) approximately 30 base pairs downstream of the

transcription site; this functions cooperatively with the Inr sequence.

I. General Transcription Factors and Transcriptional Initiation

The first step in the formation of a transcription complex is the binding of a general transcription

factor called TFIID to the promoter. TFIID is itself composed of multiple subunits, including the

TATA-binding protein (TBP) and at least 14 other polypeptides, called TBP-associated factors (TAFs).

TBP binds specifically to the TATA box while other TAF subunits of TFIID appear to bind the Inr

and DPE sequences. The binding of TFIID is followed by the binding of a second general

transcription factor, TFIIB. In addition to TBP, this factor also binds to a DNA sequence upstream of

the TATA box, known as the B recognition element (BRE). TFIIB in turn serves as a bridge to RNA

polymerase II, which binds to the TBP-TFIIB complex in association with a third factor, TFIIF.

Following recruitment of RNA polymerase II to the promoter, the binding of two additional factors

(TFIIE and TFIIH) completes formation of the initiation complex. TFIIH is a multisubunit factor that

plays at least two roles. First, XPB and XPD (two subunits of TFIIH which are also required for

nucleotide excision repair) act as helicases, unwinding DNA around the initiation site. Another subunit

of TFIIH is a protein kinase that phosphorylates tandem repeated sequences present in the C-terminal

domain (CTD) of the largest subunit of RNA polymerase II. Phosphorylation of these amino acids

releases the polymerase from its association with the preinitiation complex. Phosphorylation is further

responsible for recruitment of other proteins that allow the polymerase to initiate synthesis. The

sequential recruitment of these five general transcription factors (TFIID, TFIIB, TFIIF, TFIIE, and

TFIIH), shown assembled as the pre-initiation complex in Figure 36.1, and RNA polymerase II

represent the minimal requirements for transcription to begin in vivo.

Figure 36.1: The transcriptional pre-initiation complex (PIC) assembled along a

template DNA strand


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Within the cell, additional factors are required, including a large, multi-subunit protein complex called

Mediator, which stimulates basal transcription and plays a role in linking the general transcription

factors to the gene-specific transcription factors that regulate transcription.

II. Transcription of RNA polymerases I and III

RNA polymerase I transcribes only the genes that encode rRNA. Transcription of these genes yields a

large 45S pre-RNA, which is then processed to yield the 28S, 18S, and 5.8S rRNAs. The promoters of

ribosomal RNA genes are recognized by two transcription factors, upstream binding factor (UBF) and

selectivity factor 1 (SL1). The SL1 transcription factor is composed of four subunits, one of which is

TBP. Since the promoters of rRNA do not contain a TATA box, TBP does not bind specific promoter

sequences. Instead, the association of TBP with rRNA is mediated by the binding of other proteins in

the SL1 complex to the promoter. This is roughly analogous to the association of Inr sequences of

polymerase II genes that lack TATA boxes. Assembly of the RNA polymerase I pre-initiation complex

is shown in Figure 36.2.

Figure 36.2: Assembly of the RNA polymerase I (Pol I) pre-initiation complex

(PIC) involves the synergistic action of upstream binding factor (UBF) and

promoter selectivity factor SL1, which consists of the TATA-binding protein

(TBP) and three TBP-associated factors (TAFs). The interaction of transcription

initiation factor (TIFIA) and SL1 is essential for recruitment of Pol I.

The genes for tRNAs, 5S RNA, and some other small RNAs involved in splicing and protein transport

are transcribed by RNA polymerase III. These genes are transcribed from three distinct classes of

promoters, two of which lie within, rather than upstream of, the transcribed sequence. TFIIIA initiates

assembly of a transcription complex by binding to specific DNA sequences in the 5S rRNA promoter.

This binding is followed by the sequential binding of TFIIIC, TFIIIB, and polymerase III. This is

shown in Figure 36.3.


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Figure 36.3: Most RNA polymerase III-transcribed genes have internal promoters

within the transcribed region, which are recognized by the large, five-subunit

factor TFIIIC. In turn, TFIIIC recruits TFIIIB, which is composed of TBP and the

TAFs BRF1 and BDP1. TFIIIB recruits Pol III and assists it in initiating

transcription.

The promoters of the tRNA genes differ from those of 5S rRNA in that they do not contain the DNA

sequences recognized by TFIIIA. Instead, TFIIC binds directly to the promoter of tRNA genes,

serving to recruit TFIIIB and polymerase to form a transcription complex. Promoters of the third class

of genes transcribed by polymerase III, including genes encoding some of the snRNAs involved in

splicing, are located upstream of the transcription start site. These promoters contain a TATA box

(like those of polymerase II genes) as well a binding site for another factor called SNAP. SNAP and

TFIIIB bind cooperatively to these promoters, with TFIIIB binding directly to the TATA box. This is

mediated by TBP, a subunit of TFIIIB, which then recruits the polymerase to the transcription

complex, as shown in Figure 36.4.

Figure 36.4: The promoter of the U6 snRNA gene is located upstream of the

transcription start site. It contains a TATA box, which is recognized by the TATA-

binding protein (TBP) subunit of TFIIIB in cooperation with another factor called

SNAP.


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D. TRANSCRIPTIONAL REGULATION

The expression of eukaryotic genes is controlled primarily at the level of transcriptional initiation,

although it can also be regulated during elongation. As in bacteria, transcription in eukaryotes is

controlled by proteins that bind to specific regulatory sequences and modulate the activity of RNA

polymerase. An important difference between transcriptional regulation in eukaryotes and prokaryotes

results from the packaging of eukaryotic DNA into chromatin, limiting its availability to the

transcriptional machinery of the cell. As a result, modification of chromatin structure plays a central

role in the control of transcription in eukaryotic cells. Furthermore, a large and evolving body of

research indicates that noncoding RNAs, as well as proteins, regulate transcription in eukaryotic cells

via modifications in chromatin structure.

I. cis-Acting Regulatory Elements: Promoters and Enhancers

As discussed in chapter 35, transcription in bacteria is regulated by the binding of proteins to

cis-acting sequences, such as the lac operator, that control the transcription of adjacent genes. Similar

cis-acting sequences regulate expression in eukaryotes. Genes transcribed by RNA polymerase II have

core promoter elements, including the TATA box and the Inr sequence, that serve as specific binding

sites for general transcription factors. Other cis-acting sequences serve as binding sites for a wide

variety of regulatory factors that control the expression of individual genes. These cis-acting sequences

are frequently (although not always) located upstream of the TATA box. For example, two regulatory

elements that are found in many eukaryotic genes were first identified by studies of the promoter

region of the herpes simplex virus genes that encodes thymidine kinase. Both of these sequences are

located within 100 upstream base pairs of the TATA box. Their consensus sequences are CCAAT and

GGGCGG (called a GC box). Specific proteins that bind to these sequences and stimulate

transcription have since been identified.

In contrast to the relatively simple organization of CCAAT and GC boxes, many genes in

mammalian cells are controlled by regulatory sequences located farther from the transcription start

site. Called enhancers, these sequences were identified during studies of the promoter of another virus,

SV40. In addition to a TATA box and a set of GC boxes, two 72-base-pair repeats located farther

upstream are required for efficient transcription from this promoter. Enhancers, like promoters,

function by binding transcription factors that then regulate RNA polymerase. This is possible due to

DNA looping, which allows a transcription factor bound to a distant enhancer to interact with proteins

associated with the RNA polymerase/Mediator complex at the promoter. This is depicted in Figure

36.5.


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Figure 36.5: Transcription factors bound at distant enhancers are able to interact with the

RNA polymerase II/Mediator complex at the promoter. Because the intervening DNA

can form loops, no fundamental difference exists between the action of transcription

factors bound to DNA just upstream of the gene in the promoter and that of distant

enhancers.

Transcription factors bound to distant enhancers can thus function by the same mechanism as those

bound adjacent to promoters. Thus, there is no fundamental difference between the actions of

enhancers and those of cis-acting regulatory sequences adjacent to transcription start sites.

Enhancers can function not only over long distances, but sometimes even from different chromosomes.

This process, termed transvection, is most likely to show up on the MCAT in the context of a passage

regarding the model system in which it was first elucidated, Drosophila. Transvection is so named

because it involves trans-acting enhancers from one gene regulating the expression of the gene’s

homolog on a separate chromosome.

The binding of specific transcriptional regulatory proteins to enhancers is responsible for the

control of gene expression during development and differentiation. An example of a well-studied

enhancer is that which controls the transcription of immunoglobulins in B lymphocytes. Gene transfer

experiments have demonstrated that the immunoglobulin enhancer is active in lymphocytes, but not

in other cell types. Thus, this regulatory sequence is at least partly responsible for the tissue-specific

expression of the immunoglobulin genes in the appropriate cell type.

Importantly, enhancers usually contain multiple functional sequence elements that bind

different transcriptional regulatory proteins. These proteins work together to regulate gene expression.

Returning to the previous example, the immunoglobulin heavy-chain enhancer spans more than 200

base pairs and contains at least nine distinct sequence elements that serve as protein-binding sites. The


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mutation of any of these sequences reduces, but does not abolish, enhancer activity, indicating that the

functions of individual proteins that bind to the enhancer are at least partially redundant. In non-

lymphoid cells, many of the individual sequence elements of the immunoglobulin enhancer are able to

stimulate transcription by themselves. The restricted activity of the intact enhancer in B lymphocytes

therefore does not arise from tissue-specific functions of each of its components. Instead, tissue-specific

expression results from the combination of the individual sequence elements that make up the

complete enhancer. These elements include some cis-acting regulatory sequences that bind

transcriptional activators that are expressed specifically in B lymphocytes, as well as other regulatory

sequences that bind repressors in non-lymphoid cells. Accordingly, the immunoglobulin enhancer

contains negative regulatory elements that inhibit transcription in inappropriate cell types, as well as

positive regulatory elements that activate transcription in B lymphocytes. The overall activity of the

enhancer is greater than the sum of its parts, reflecting the combined action of the proteins associated

with each of the individual sequence elements.

Although DNA looping allows enhancers to act over a considerable distance from the

promoters, the activity of any given enhancer is specific for the promoter of its appropriate target gene.

This specificity is maintained in part by insulators or barrier elements, which divide chromosomes into

independent domains and prevent enhancers from acting on promoters located in an adjacent

domain. Insulators also prevent the chromatin structure of one domain from expanding into the

region occupied by a neighboring chromatin structure, thereby maintaining independently regulated

regions of the genome. It is thought that insulators function separately by organizing independent

domains of chromatin within the nucleus, but their mechanism of action remains a subject of ongoing

research. One potential application of insulator elements relates to gene therapy, where a major

hurdle is preventing the aberrant regulation or inactivation of an introduced gene by the nearby

chromatin structure. The addition of an insulator element is a potential solution to this problem.

II. Transcription Factor Binding Sites

The binding sites of transcriptional regulatory proteins in promoter or enhancer sequences have

commonly been identified by two types of experiments. The first, footprinting, was originally

developed to characterize the binding of RNA polymerase to prokaryotic promoters. In experiments of

this type, shown in Figure 36.6, a DNA fragment is radiolabeled at one end. The labeled DNA is

incubated with the protein of interest (e.g., RNA polymerase) and then subjected to partial digestion

with DNase.


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Figure 36.6: DNA footprinting. 1. A sample containing fragments of DNA is radiolabeled

at once end. 2. The sample is then divided in two. Half is incubated with a protein that

binds to specific sequences of DNA within the fragment, while the remainder is

unchanged. 3. Both samples are then digested with DNAase, under conditions such that

the DNAase introduces an average of one cut per molecule. The region of DNA bound to

the protein is protected from DNAase digestion. The DNA-protein complexes are then

denatured, and the sizes of the radiolabeled DNA fragments produced by DNAase

digestion are analyzed by electrophoresis. Fragments of DNA resulting from DNAase

cleavage within the region protected by protein binding are missing from the sample of

DNA that was incubated with protein.

The principle of the method is that the regions of DNA to which the protein binds are protected from

DNase digestion. These regions can therefore be identified by comparison of the digestion products of

the protein-bound DNA with those resulting from identical DNase treatment of a parallel sample of

DNA that was not incubated with proteins. Variations of this method, which employ chemical

reagents that modify and cleave DNA at particular nucleotides, can be used to identify the specific

DNA bases that are in contact with protein.

A second approach involves performing an electrophoretic-mobility shift assay in which a

radiolabelled DNA fragment is incubated with a protein preparation and then subjected to

electrophoresis through a nondenaturing gel. Protein binding is indicated by a decrease in the


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electrophoretic mobility of the DNA fragment, since the bound protein slows its migration through the

gel. The combined use of footprinting and electrophoretic mobility shifts has led to the correlation of

protein-binding sites with the regulatory elements of enhancers and promoters, indicating that these

sequences generally constitute the recognition sites of specific DNA-binding proteins.

The binding sites of most transcription factors consist of short, degenerate DNA sequences,

meaning that the transcription factor will bind not only to the consensus sequence but also to

sequences that differ from the consensus at one or more positions. Because of their short, degenerate

nature, sequences matching transcription factors occur frequently in genomic DNA, so physiologically

significant regulatory sequences cannot be identified using DNA sequences alone. Such identification

remains one of the primary challenges in molecular biology. One experimental approach is that of

chromatin immunoprecipitation (ChIP). Cells are first treated with formaldehyde, which cross-links

proteins to DNA. As a result, transcription factors are covalently linked to the DNA sequences to

which they were bound within the living cell. Chromatin is then extracted and sheared to fragments of

about 500 base pairs. Fragments of DNA linked to a transcription factor of interest can then be

isolated by immunoprecipitation with an antibody against the transcription factor. The formaldehyde

crosslinks are then reversed, and the immunoprecipitated DNA is isolated and analyzed to determine

the sites to which the specific transcription factor was bound within the cell. A ChIP sequencing

protocol is shown in Figure 36.7.


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Figure 36.7: Chromatin immunoprecipitation (ChIP). 1. Sample cells are treated with a

reversible chromatin cross-linking agent. 2. DNA strands are sheared by sonication. 3.

Bead-attached antibodies are added to immunoprecipitate target transcription factor

protein. 4. DNA is unlinked, yielding purified DNA. 5. Purified DNA extract is

sequenced.


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III. Transcriptional Regulatory Proteins

A wide range of transcriptional regulatory proteins have been isolated and identified based on their

specific binding affinity for select DNA sequences. Because transcription factors are central to the

regulation of gene expression in eukaryotes, they remain a major area of ongoing research in cellular

and molecular biology, and a frequently tested molecular genetics topic on the MCAT. Of these

proteins, the best understood are transcriptional activators, which bind to regulatory DNA sequences

and stimulate transcription. In general, these factors consist of two independent domains; one region

specifically binds to DNA, while the other stimulates transcription by interacting with other proteins,

including Mediator or other components of a cell’s transcriptional machinery. The basic function of

the DNA-binding domain is to anchor the transcription factor to the proper DNA site. The activation

domain then independently stimulates transcription through protein-protein interaction.

IV. Transcription Factors

More than 2500 transcription factors encoded by the human genome have thus far been recognized.

They contain a diversity of distinct DNA-binding domains, the most common of which is the zinc

finger domain. This type of domain consists of repeating cysteine and histidine residues that bind zinc

ions and fold into DNA-binding loop structures referred to as “fingers,” the structure of which is

shown in Figure 36.8.


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Figure 36.8: Structural representation of the Cys2His2 zinc finger motif, consisting

of an α helix and an antiparallel β sheet. The zinc ion (center) is coordinated by

two histidine residues and two cysteine residues.

These domains were initially identified in the polymerase III transcription factor TFIIIA, but they are

also common among transcription factors that regulate polymerase II promoters, including Sp1, one

of the earliest identified and most common eukaryotic transcription factors. Other examples of

transcription factors that contain zinc finger domains are the steroid hormone receptors, which

regulate gene transcription in response to hormones such as estrogen and testosterone.

Artificial transcription factors with zinc-finger domains designed to bind specific sequences

within the genome have been developed. By ligating different effector domains to the DNA binding

domain, the target gene can be either activated or repressed. This technology can be applied to gene

therapy and the development of transgenic plants and animals of commercial interest.

The helix-turn-helix motif (Figure 36.9) was first recognized in prokaryotic DNA-binding

proteins, including the E. coli catabolite activator protein (CAP).

Figure 36.9: Basic helix-turn-helix structural motif. Two α-helices are connected by a

short loop.

In these proteins, one of the helices makes most of the contact with DNA, while the other lies across

the complex in order to stabilize the interaction. In eukaryotes, helix-turn-helix proteins include the

homeodomain proteins, which play essential roles in the regulation of gene expression during

embryonic development. Genes encoding these proteins were first identified in developmental mutants

of Drosophila. Some of the earliest recognized Drosophila mutants resulted in the development of flies in


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which one body part was transformed into another. For example, in one homeotic mutant, legs rather

than antennae grow from the head of the fly. Genetic analysis has shown that these mutants contain

nine homeotic genes, each of which specifies a different body segment. Molecular cloning and analysis

of these genes indicate that they contain conserved sequences of 180 base pairs, called homeoboxes,

which encode DNA-binding domains (homeodomains) of transcription factors. A wide variety of

similar homeodomain proteins have since been identified in fungi, plants and other animals, including

humans.

Two other families of DNA-binding proteins – the leucine zipper and helix-loop-helix

proteins – contain DNA-binding domains formed by the dimerization of two polypeptide chains. The

leucine zipper contains four or five leucine residues spaced at intervals of seven amino acids, resulting

in their hydrophobic side chains being exposed at one side of a helical region. This region serves as the

dimerization domain for the two protein subunits, which are held together by hydrophobic

interactions between the leucine side chains. Immediately following the leucine zipper is a region rich

in positively charged lysine and arginine residues that bind DNA. This interaction is depicted in Figure

36.10.

Figure 36.10: Leucine zipper dimer bound to DNA fragment

The helix-loop-helix proteins are similar in structure, but differ in the exact structure of their

dimerization domains, formed by two helical regions separated by a loop. An important feature of

both leucine zipper and helix-loop-helix transcription factors is that different members of each family

can dimerize with one another. Thus, the combination of distinct protein subunits can form an

expanded array of factors that can differ in both DNA sequence recognition and transcription-


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stimulating activity. This formation of dimers between different family members is a critical aspect of

their self-regulation. Both families of DNA-binding proteins play a central role in regulating tissue-

specific and inducible gene expression.

The activation domains of transcription factors are not as well characterized as their DNA-

binding domains. Some, called acidic activation domains, are rich in negatively charged residues such

as aspartate and glutamate; others are rich in proline or glutamine. The activation domains appear to

stimulate transcription by two distinct mechanisms. First, they interact with Mediator proteins and

general transcription factors, such as TFIIB or TFIID, to recruit RNA polymerase; in doing so, they

facilitate the assembly of a transcription complex on the promoter, similar to that which occurs in

transcriptional activators in bacteria. In addition, eukaryotic transcription factors interact with a

variety of coactivators that stimulate transcription by modifying chromatin structure, as discussed later

in this chapter.

Figure 36.11: Eukaryotic activators stimulate transcription by exerting two downstream

effects: 1) they interact with Mediator proteins and general transcription factors to

facilitate the assembly of a transcription complex and stimulate transcription, and 2) they

interact with coactivators that facilitate transcription by modifying chromatin structure.

V. Eukaryotic Repressors

Eukaryotic gene expression is regulated not just by transcriptional activators, but also by

repressors. Like their prokaryotic counterparts, eukaryotic repressors bind to specific DNA sequences

and inhibit transcription. In some cases, eukaryotic repressors act by interfering with the binding of

other transcription factors to DNA. For example, the binding of a repressor near the transcription start

site can block the interaction of RNA polymerase or general transcription factors with the promoter,


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an action similar to that of bacterial repressors. Other repressors compete with activators for binding

to specific regulatory sequences. Some of these repressors contain the same DNA-binding domain as

the activator, but lack its activation domain. As a result, their binding to a promoter or enhancer

blocks the binding of the activator, inhibiting transcription. This is shown in Figure 36.12.

Figure 36.12: Some repressors block the binding site of activators to regulatory

sequences.

In contrast to repressors that simply interfere with activator binding, “active” repressors

contain specific functional domains that inhibit transcription via protein-protein interaction. Many

active repressors have been shown to play key roles in the regulation of transcription related to

proteins that control cellular growth and differentiation. As with transcriptional activators, several

distinct types of repressor domains have been identified. For example, the repressor domain of one of

the first eukaryotic repressor proteins to be identified, Krüppel, is rich in alanine residues, while other

repression domains are rich in proline or acidic residues. The functional targets of repressors are

equally diverse. Repressors can inhibit transcription by interacting with specific activator proteins,

with Mediator proteins, with general transcription factors, or with corepressors that act by modifying

chromatin structure. An example of one such mode of repression is seen in Figure 36.13.


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Figure 36.13: Some repressors have active repression domains that inhibit transcription

by interactions with Mediator proteins or general transcription factors, as well as with co-

repressors that act to modify chromatin struture.

The regulation of transcription by both repressors and activators considerably extends the

range of mechanisms that control the expression of eukaryotic genes. One important role of repressors

may be to inhibit the expression of tissue-specific genes in inappropriate cell types. For example, as

noted earlier, a repressor binding site in the immunoglobulin enhancer is thought to contribute to its

tissue-specific expression by suppressing transcription in non-lymphoid cell types. Other repressors

play important roles in the control of cell proliferation and differentiation in response to signaling by

hormones and growth factors.

E. REGULATION OF CHROMATIN STRUCTURE

As referenced in the preceding discussion, both activators and repressors regulate transcription in

eukaryotes not only by interacting with Mediator and other components of the transcriptional

machinery, but also by inducing changes in the structure of chromatin. Rather than being present

within the nucleus as naked genetic material, the DNA of all eukaryotic cells is tightly bound to

histones. The basic structural unit of chromatin is the nucleosome, each of which consists of 147 base

pairs of DNA wrapped around the core histones H2A, H2B, H3, and H4. The core histones are

present in the nucleosome as an octamer containing four dimers, with one molecule of histone H1

bound to the DNA as it enters the nucleosome core particle. The organization of the nucleosome is

shown in Figure 36.14 and the formation of a histone core octamer is shown in Figure 36.15.


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Figure 36.14: Nucleosome organization

Figure 36.15: Formation of the histone octamer

The chromatin is further condensed by being coiled into higher-order structures organized into large

loops of DNA. This packing of eukaryotic DNA in chromatin has significant consequences in terms of

the packaged DNA’s availability as a template for transcription; control over the state in which cellular

DNA is maintained represents an important regulatory tool in eukaryotic cells.

Actively transcribed genes are found in relatively de-condensed chromatin, roughly

corresponding to 30-nm fibers. Nonetheless, actively transcribed genes remain bound in a relatively


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inaccessible state to histones and packaged in nucleosomes, presenting transcription factors and RNA

polymerase with the problem of interacting with DNA in a nucleosome structure. The tight winding of

DNA around the nucleosome core particle is a major obstacle to transcription, affecting both the

ability of transcription factors to bind DNA and the ability of RNA polymerase to transcribe through

the complex spatial arrangement of a chromatin template.

I. Histone Modification

Several modifications are characteristic of transcriptionally active chromatin, including histone

modifications, nucleosome rearrangements, and the association of two non-histone chromosomal

proteins, called HMGN proteins, with the nucleosomes of actively transcribed genes. The binding sites

of the HMGN proteins on nucleosomes overlap the binding site of histone H1, and it appears that

these proteins stimulate transcription by affecting modifications of histone H1 to maintain a

decondensed chromatin structure.

Figure 36.16: The binding of epigenetic factors to histone “tails” alters the extent to

which DNA is wrapped around histones and the availability of genes in the DNA to be

activated.

Histone acetylation has been correlated with transcriptionally active chromatin in a wide

variety of cell types. The core histones have two domains: a histone fold domain, which is involved in

the interaction with other histones and in wrapping DNA around the nucleosome core particle, and an

amino-terminal domain, which extends outside the nucleosome. The amino-terminal domains are rich

in lysine and can be modified by acetylation at specific lysine residues. Histone acetyltransferases have

been associated with a number of mammalian transcriptional coactivators, as well as with general


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transcription factor TFIID. Conversely, transcriptional corepressors in both yeast and mammalian

cells function as histone deacetylases, which remove acetyl group from histone tails. Histone

acetylation is thus targeted directly by both transcriptional activators and repressors, indicating that it

plays a role in regulation of eukaryotic gene expression.

Histones are modified not only by acetylation, but also by phosphorylation of serine residues,

methylation of lysine and arginine residues, and addition of ubiquitin to lysine residues. Like

acetylation, these modifications occur at specific amino acid residues in the histone tails that are

associated with changes in transcriptional activity. Changes in gene expression brought about by

histone modification are likely a result of the creation of binding sites for other regulatory proteins.

According to this hypothesis, combinations of histone modifications constitute a “histone code” that

regulates gene expression by recruiting other regulatory proteins to the chromatin template. For

example, transcriptionally active chromatin is associated with several specific modifications of histone

H3; these include methylation of lysine-4, phosphorylation of serine-10, acetylation of lysine-9, -14, -

18, and -23, and methylation of arginine-17 and -26. In contrast, methylation of H3 lysine-9 leads to

the suppression of target genes by the recruitment of corepressors. The methylated H3 lysine-9

residues have further been shown to serve as binding sites for proteins that induce chromatin

condensation, directly linking the histone modification to transcriptional repression and the formation

of heterochromatin. Additionally, these modifications of histone tails may also regulate one another,

leading to the establishment of distinct patterns of histone modification that correlate with stable

modification in transcriptional activity.


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Figure 36.17: Histone modifications play fundamental roles in most biological processes

that are involved in the manipulation and expression of DNA. In multicellular organisms,

facultative heterochromatin regions contain genes that are differentially expressed

through development and/or differentiation and which then become silenced while

constitutive heterochromatin contains permanently silenced genes in genomic regions

such as the centromeres and telomeres. Euchromatin is a far more relaxed environment

containing active genes.

II. Nucleosome Remodeling Factors

In contrast to the enzymes that regulate chromatin structure by modifying histones, nucleosome

remodeling factors are protein complexes that alter the arrangement or structure of nucleosomes. One

of their mechanisms of action involves catalyzing the sliding movement of histone octamers along the

DNA molecules, thereby repositioning the nucleosomes to change the transcription factor accessibility

of specific DNA sequences. Alternatively, nucleosome remodeling factors may act by inducing changes

in the conformation of nucleosomes, again affecting the ability of specific DNA sequences to interact


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with transcriptional regulatory proteins. Like histone modifying enzymes, nucleosome remodeling

factors can be recruited to DNA in association with either transcriptional activators or repressors, and

can alter the arrangement of nucleosomes to either stimulate or inhibit transcription.

III. Regulation of Transcriptional Elongation

The recruitment of histone modifying enzymes and nucleosome remodeling factors by transcriptional

activators stimulates the initiation of transcription by altering the chromatin structure of enhancer and

promoter regions. However, following initiation, RNA polymerase must still elongate the nascent

mRNA transcript through the structurally complex chromatin template. This is facilitated by

elongation factors that become associated with the phosphorylated C-terminal domain of RNA

polymerase II at the initiation of transcription. These elongation factors include histone modifying

enzymes (acetyltransferases and methyltransferases) as well as proteins that transiently disrupt the

structure of nucleosomes during transcription. Although it has been less thoroughly studied than

transcriptional initiation (and is therefore a less likely MCAT passage topic), transcriptional elongation

does present an additional level at which gene expression can be regulated in eukaryotic cells.

IV. DNA Methylation

The methylation of DNA is another general mechanism that controls transcription in eukaryotes.

Cytosine residues in the DNA of fungi, plants, and animals can be modified by the addition of methyl

groups at the 5-carbon position. DNA is methylated specifically at the cytosine residues that precede

guanines in the DNA chain, known as CpG dinucleotides. This methylation is correlated with

transcriptional repression. Methylation commonly occurs within transposable elements, where it

appears that methylation suppresses the movement of transposons throughout the genome. In

addition, DNA methylation is associated with transcriptional repression of some genes along with

alterations in chromatin structure. In plants, miRNAs direct DNA methylation as well as chromatin

modifications of repressed genes; it is unclear whether this also occurs in animals. However, it is

known that genes on the inactive X chromosomes in mammals become methylated following

transcriptional repression by Xist RNA. It appears true, then, that DNA methylation, as well as histone

modification, plays a role in X chromosome inactivation.

One important regulatory role of DNA methylation has been established in the phenomenon

known as genomic imprinting, which controls the expression of some genes involved in mammalian

embryonic development. In most cases, both the maternal and paternal alleles are expressed in diploid

cells. However, there are some imprinted genes which show varied expression depending on whether

they are inherited from the mother or from the father. In some cases, only the paternal allele of an


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imprinted gene is expressed, and the maternal allele is transcriptionally inactive. For other imprinted

genes, the maternal allele is expressed and the paternal allele is inactive.

DNA methylation appears to play an important role in distinguishing between the paternal

and maternal alleles of imprinted genes. A good example is the gene H19, which is transcribed only

from the maternal copy. The H19 gene is specifically methylated during the development of male, but

not female, germ cells. The union of sperm and egg at fertilization therefore yields an embryo

containing a methylated paternal allele and an unmethylated maternal allele of the gene. Following

DNA replication, these differences are maintained by an enzyme that specifically methylates CpG

sequences of a daughter strand that is hydrogen-bonded to a methylated parental strand. As a result,

the paternal H19 allele remains methylated, staying transcriptionally inactive in both embryonic and

somatic tissues. The paternal H19 allele does become demethylated in the germ line, however,

allowing a new pattern of methylation to be established for transmission to the next generation.

F. ROLE OF NONCODING RNA

Recent advances indicate that gene expression can be regulated not only by the transcriptional

regulatory proteins already discussed, but also by noncoding RNA molecules. One mode of action of

noncoding regulatory RNAs is to inhibit translation by RNA interference, a phenomenon in which

short double-stranded RNAs induce degradation of a homologous mRNA. In addition, noncoding

RNAs can repress transcription by inducing histone modifications that lead to chromatin condensation

and the formation of heterochromatin. MicroRNAs (miRNAs) are naturally-occurring short

noncoding RNAs that function as normal regulators of gene expression. Hundreds of genes encode

miRNAs in both plants and animals, so it appears that gene regulation by these noncoding RNAs is a

widespread phenomenon, even though the functions of most miRNAs have yet to be determined.

miRNAs are transcribed as precursors containing inverted stem-loop structures. These

precursors are then cleaved by an enzyme known as Dicer to form mature miRNAs, which are short

double-stranded RNAs of approximately 20-25 nucleotides. In RNA interference, miRNAs associate

with the RNA-induced silencing complex (RISC), within which the two strands of miRNA separate

and target homologous mRNAs for cleavage. This is shown in Figure 36.18.


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Figure 36.18: RNA interference. 1. Stem-loop structure forms via hydrogen bonding. 2.

Dicer-catalyzed cleavage yields short, mature miRNA segments. 3. One strand of each

segment is degraded; the other strand associates with the protein complex, RISC. 4. The

miRNA-bound complex can base-pair with any target mRNA that contains a sufficiently

complementary sequence. 5. The miRNA-protein complex prevents gene expression

either by degrading target mRNA or blocking its translation.

In transcriptional repression, the miRNAs associate with a different protein complex called the RITS

(RNA-induced transcriptional silencing) complex. The separated miRNA strands then guide the RITS

complex to the homologous gene, most likely by base pairing with the mRNA transcript in association

with RNA polymerase II. RITS then represses transcription by recruiting a histone methyltransferase

that methylates histone H3 lysine-9, leading to the formation of transcriptionally inactive

heterochromatin.

I. X Chromosome Inactivation

The phenomenon of X chromosome inactivation provides another example of the role of noncoding

RNA in regulating gene expression in mammals. In many animals, including humans, females have

two X chromosomes while males have one X and one Y chromosome. The X chromosome contains

hundreds of genes that are not present on the much smaller Y chromosome. Thus, females have twice

the number of X chromosome genes found in most males. Despite this difference, female and male

cells contain equal amounts of the proteins encoded by the majority of X chromosome genes. This

results from a dosage compensation mechanism in which the large majority of genes on one of the two

X chromosomes in female cells are inactivated by being converted to heterochromatin early in


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development. Consequently, only one copy of most genes located on the X chromosome is available

for transcription in the cells of either females or males.

While the mechanism of X chromosome inactivation has not yet been fully elucidated, the key

elements appear to be a noncoding RNA transcribed from a regulatory gene, called Xist, on the

inactive X chromosome. Xist RNA remains localized to the inactive X, binding to and coating the

chromosome. This leads to the recruitment of a protein complex that induces methylation of histone

H3 lysine-27 and lysine-9, leading to chromatin condensation and conversion of most of the X genes

to heterochromatin.

G. POST-TRANSCRIPTIONAL CONTROL AND RNA PROCESSING

Although transcription is the first and most highly regulated step in gene expression, it is usually only

the beginning of the series of events required to produce functional RNA. Most newly synthesized

RNAs must be modified in various ways to be converted to their functional forms. Bacterial mRNAs

are an exception; they are used immediately as templates for protein synthesis while still being

transcribed. However, the primary transcripts of both rRNAs and tRNAs must undergo a series of

processing steps in prokaryotic as well as eukaryotic cells. Primary transcripts of eukaryotic mRNAs

similarly undergo extensive modifications, including the removal of introns by splicing, before they are

transported from the nucleus to the cytoplasm to serve as templates for protein synthesis. Regulation of

these processing steps and regulation of the rate of mRNA degradation in the cell provide another

level of control of gene expression.

I. Processing of Ribosomal and Transfer RNA

The basic processing of rRNA and tRNA in prokaryotes and eukaryotes is similar, as might be

expected given the common roles of these molecules in protein synthesis. As discussed already,

eukaryotes have four types (5S, 5.8S, 18S, 28S) of ribosomal RNAs, three of which (28S, 18S, and 5.8S

rRNAs) are derived by cleavage of a single long precursor transcript, called pre-rRNA. Prokaryotes

have three ribosomal rRNAs (5S, 23S, 23S), which are equivalent to the 28S, 18S and 5S rRNAs of

eukaryotic cells and are also formed by the processing of a single pre-rRNA transcript. The only

rRNA that is not processed extensively is the 5S rRNA, which is transcribed from a separate gene.

I n eukaryotic cells, processing of rRNA takes place within the nucleolus of cells. During

processing, pre-rRNA is first cleaved at a site adjacent to the 5.8S rRNA on its 5’ side, yielding two

separate precursors that contain the 18S and 28S + 5.8S rRNAs, respectively. Further cleavage then

converts these to their final products, with the 5.8S rRNA becoming hydrogen-bonded to the 28S

molecule. In addition to these cleavages, rRNA processing involves the addition of methyl groups to


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bases and sugar moieties of specific nucleotides and the conversion of some uridine residues to

pseudouridine.

Like rRNAs, tRNAs in both bacteria and eukaryotes are synthesized as longer precursor

molecules known as pre-tRNAs, some of which contain several individual tRNA sequences. The

processing of the 5’ end of pre-tRNA involves cleavage by the enzyme RNase P; this reaction is of

particular interest, as it is the prototypical model of catalysis by an RNA enzyme. RNase P consists of

RNA and protein components, both of which are required for maximal activity; however, the catalytic

activity of RNase P is due to its RNA component. For this reason, RNase P is categorized as a

ribozyme.

The 3’ end of tRNA is generated by the action of a protein RNase, but the processing of this

end of the tRNA molecule also involves an unusual activity: the addition of a CCA terminus. All

tRNAs have the sequence CCA at their 3’ ends. This sequence is the site of amino acid attachment, so

it is required for tRNA function during protein synthesis. The CCA terminus is encoded in the DNA

of some tRNA genes; in others, it is instead added as an RNA processing step by an enzyme that

recognizes and adds CCA to the 3’ end of all tRNAs that lack this sequence.

Another unusual characteristic of tRNA processing is the extensive post-transcriptional

modification of bases in tRNA molecules. Approximately 10% of all bases in tRNAs are altered to

yield a variety of modified nucleotides at specific positions. The functions of most of these modified

bases are unknown, but some play important roles in protein synthesis by altering the base-pairing

properties of tRNA molecules.

Some pre-tRNAs, as well as pre-rRNA in a few organisms, contain introns that are removed

by splicing. In contrast to other splicing reactions, which involve the activities of catalytic RNAs,

tRNA splicing is mediated by conventional protein enzymes. An endonuclease cleaves the pre-tRNA

at the splice sites to excise the intron, followed by joining of the exons to form a mature tRNA

molecule.

II. Processing of mRNA in Eukaryotes

In contrast to the processing of rRNA and tRNA, the ways in which mRNA is processed by eukaryotes

and prokaryotes is substantially different. In bacteria, ribosomes have immediate access to mRNA,

allowing translation to begin on the nascent mRNA chain while transcription is still underway. In

eukaryotes, mRNA synthesized in the nucleus must first be transported to the cytoplasm before it can

be used as a template for protein synthesis. Moreover, the initial products of transcription in

eukaryotic cells, called pre-mRNAs, are extensively modified before export from the nucleus. The

processing of mRNA includes modification of both ends of the initial transcript, as well as the removal

of introns. Rather than this occurring as a series of independent events following synthesis of pre-


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mRNA, these processing reactions are closely coordinated steps in gene expression. The C-terminal

domain (CTD) of RNA polymerase II plays a key role in coordinating these processes by serving as a

binding site for the enzyme complexes involved in mRNA processing. The association of these

processing enzymes with the CTD of RNA polymerase II accounts for their specificity in processing

mRNAs; RNA polymerases I and III lack a CTD, so their transcripts are not processed by the same

enzyme complexes.

The first step in mRNA processing is the modification of the 5’ end of the transcript by the

addition of a structure called a 7-methylguanosine cap. The enzymes responsible for capping are

recruited to the phosphorylated CTD following initiation of transcription, and the cap is added after

transcription of the first 20-30 nucleotides of the RNA. Capping is initiated by the addition of a GTP

in reverse orientation to the 5’ terminal nucleotide of the RNA. Methyl groups are then added to this

guanosine residue and to the ribose moieties of one or two 5’ nucleotides of the RNA chain. The 5’

cap stabilizes the RNA and aligns eukaryotic mRNAs on the ribosome during translation.

The 3’ end of most eukaryotic mRNAs undergoes modification as well. This end of the

molecule is defined not by termination of transcription, but by cleavage of the primary transcript and

addition of a poly-A-tail via a processing reaction called polyadenylation. The signals for

polyadenylation include a highly conserved hexanucleotide located 10 to 30 nucleotides upstream of

the site of polyadenylation; its sequence is AAUAAA in mammalian cells. A G-U rich downstream

sequence element acts as another signal. In addition, some genes have a U-rich sequence element

upstream of the AAUAAA. These sequences are recognized by a complex of proteins, including an

endonuclease that cleaves the RNA chain and a separate poly-A polymerase that adds a poly-A tail of

about 200 nucleotides to the transcript. These processing enzymes are associated with the

phosphorylated CTD of RNA polymerase II, and may travel with the polymerase, beginning at the

transcription initiation site. Cleavage and polyadenylation is followed by degradation of the RNA that

has been synthesized downstream of the site of poly-A addition, resulting in the termination of

transcription.

Almost all mRNAs in eukaryotes are polyadenylated, and poly-A tails have been shown to

regulate both translation and mRNA stability. In addition, polyadenylation plays an important

regulatory role in early development, where changes in the length of poly-A tails control mRNA

translation. For example, many mRNAs are stored in unfertilized eggs in an untranslated form with

short poly-A tails. Fertilization stimulates the lengthening of the poly-A tails of these stored mRNAs,

which in turn activates their translation and the synthesis of proteins required for early embryonic

development.

The most structurally dramatic modification of pre-mRNA is the removal of introns by

splicing. The coding sequences of most eukaryotic genes are interrupted by noncoding sequences

(introns) that are precisely excised from the mature mRNA. In mammals, most genes contain multiple


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introns, which typically account for about ten times more pre-mRNA sequences than do the exons.

Studies of the mechanism of splicing has illuminated new mechanisms of gene regulation, and has

revealed novel catalytic activities of RNA molecules.

Figure 36.19: Formation of the primary transcript and its processing in a eukaryotic cell

in the nucleus. The 5’ cap is added before synthesis of the primary transcript is complete.

A noncoding sequence (intron) following the last exon is shown. Splicing can occur

either before or after cleavage and polyadenylation.

H. SPLICING MECHANISMS

The key to understanding pre-mRNA splicing was the development of in vitro systems that efficiently

carried out the splicing reaction. Pre-mRNAs were synthesized in vitro by the cloning of structural

genes, including their introns, adjacent to promoters for bacteriophage RNA polymerases, which

could readily be isolated in large quantities. Transcription of these plasmids could then be used to

prepare large amounts of pre-mRNAs that, when added to nuclear extracts of mammalian cells, were

found to be correctly spliced. As with transcription, the use of such in vitro systems has allowed splicing

to be analyzed in much greater detail than would have been possible in intact cells.

Analysis of the reaction products and intermediates formed revealed that pre-mRNA splicing

proceeds in two steps. First, pre-mRNA is cleaved at the 5’ splice site, and the 5’ end of the intron is

joined to an adenine nucleotide within the intron (near its 3’ end). In this step, an unusual bond is

formed between the 5’ end of the intron and the 2’ hydroxyl group of the adenine nucleotide. The

resulting intermediate is a lariat-like structure in which the intron forms a loop. The second step in


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splicing then proceeds with simultaneous cleavage at the 3’ splice site and ligation of the two exons.

The intron is thus excised as this lariat-like structure, which is then linearized and degraded within the

nucleus of the intact cells.

These reactions define three critical sequence elements of pre-mRNAs: sequences at the 5’

splice site, sequences at the 3’ splice site, and sequences within the intron at the branch point (the point

at which the 5’ end of the intron becomes ligated to form the lariat-like structure). Pre-mRNAs contain

similar consensus sequences at each of these positions, allowing the splicing apparatus to recognize

pre-mRNAs and carry out the cleavage and ligation reactions involved in the splicing process.

Biochemical analysis of nuclear extracts has revealed that splicing takes place in large

complexes, called spliceosomes, composed of proteins and RNAs. The RNA components of the

spliceosome are five types of small nuclear RNAs (snRNAs) called U1, U2, U4, U5, and U6. These

snRNAs, which range in size from approximately 50 to nearly 200 nucleotides, are complexed with six

to ten protein molecules to form small nuclear ribonucleoprotein particles (snRNPs), which play

central roles in the splicing process. The U1, U2, and U5 snRNPs each contain a single snRNA

molecule, whereas U4 and U6 snRNAs are complexed to each other in a single snRNP.

The first step in spliceosome assembly involves the binding of U1 snRNP to the 5’ splice site

of pre-mRNA. This recognition of 5’ splice sites involves base pairing between the 5’ splice site

consensus sequence and a complementary sequence at the 5’ end of U1 snRNA. U2 snRNP then

binds to the branch point using similar complementary base pairing between U2 snRNA and branch

point sequences. A preformed complex consisting of U4/U6 and U5 snRNPs is then incorporated into

the spliceosome, with U5 binding to sequences upstream of the 5’ splice site. The splicing reaction is

then accompanied by rearrangements of the snRNAs. Prior to the first reaction steps (leading to the

formation of the lariat-like intermediate), U6 dissociates from U4 and displaces U1 at the 5’ splice site.

U5 then binds to sequences at the 3’ splice site, followed by excision of the intron and ligation of the

exons.

Not only do the snRNAs recognize consensus sequences at the branch points and splice sites

of pre-mRNAs, but they also directly catalyze the splicing reaction. The catalytic role of RNAs in

splicing was demonstrated by the discovery that some RNAs are capable of self-splicing; that is, they

can catalyze the removal of their own introns in the absence of other proteins or RNA factors. Self-

splicing was first described during studies of 28S rRNA from protozoa. Further studies have shown

that splicing is catalyzed by the intron, which acts as a ribozyme to direct its own excision from the

pre-rRNA molecule. Additional studies have revealed self-splicing RNAs in mitochondria,

chloroplasts, and bacteria. These self-splicing RNAs are divided into two classes on the basis of their

reaction mechanisms: group I and group II introns. The first step in the splicing of group I introns is

cleavage at the 5’ splice site mediated by guanosine cofactor. The 3’ end of the free exon then reacts


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with the 3’ splice site to excise the introns as a linear RNA molecule. The splicing mechanism of group

I introns is illustrated in Figure 36.20.

Figure 36.20: Splicing mechanism of group I introns. 1. The 3’ OH group of guanosine,

GMP, GDP or GTP attacks the phosphate located at the 5’ splice site. 2. The 3’ OH of the

5’ exon becomes the nucleophile, completing the reaction.

In contrast, the self-splicing reactions of group II introns (as found in some mitochondrial pre-mRNAs)

closely resemble nuclear pre-mRNA splicing in which cleavage of the 5’ splice site results from the


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attack of an adenosine nucleotide in the intron. As with pre-mRNA splicing, the result is a lariat-like

intermediate, which is then excised.

The similarity between spliceosome-mediated pre-mRNA splicing and self-splicing of group II

introns strongly suggests that the active catalytic components of the spliceosomes are RNAs rather

than proteins. In particular, these similarities suggest that pre-mRNA splicing is catalyzed by the

snRNAs of the spliceosome. Continuing studies of pre-mRNA splicing have provided clear support of

this view, including the demonstration that U2 and U6 snRNAs, in the absence of proteins, can

catalyze the first step in pre-mRNA splicing. Pre-mRNA splicing is thus considered to be an RNA-

based reaction, catalyzed by spliceosome snNRAs acting analogously to group II self-splicing introns.

Within the cell, protein components of the snRNPs are also required, however, and participate in both

assembly of the spliceosome and the splicing reaction itself.

A number of protein splicing factors that are not snRNP components also play critical roles in

spliceosome assembly, particularly in identification of the correct splice sites in pre-mRNAs.

Mammalian pre-mRNAs typically contain multiple short exons separated by much larger introns.

Introns frequently contain sequences that resemble splice sites, so the splicing machinery must be able

to identify the appropriate 5’ and 3’ sites at intron/exon boundaries to produce a functional mRNA

molecule. Splicing factors serve to direct spliceosomes to the correct splice sites by binding to specific

RNA sequences and then recruiting U1 and U2 snRNPs to the appropriate sites on pre-mRNA by

protein-protein interactions. For example, the SR splicing factors bind to specific sequences within

exons and act to recruit U1 snRNP to the 5’ splice site. SR proteins also interact with another splicing

factor (USAF), which binds to pyrimidine-rich sequences at 3’ splice sites and recruit U2 snRNP to the

branch point. In addition to recruiting the components of the spliceosome to the pre-mRNA, splicing

factors couple splicing to transcription by associating with the phosphorylated CTD of RNA

polymerase II. This anchoring of the splicing machinery to RNA polymerase is thought to be

important in ensuring that exons are joined in the correct order as the pre-mRNA is synthesized.

I. Alternative Splicing

The central role of splicing in the processing of pre-mRNA opens the possibility of regulation of gene

expression by control of the splicing machinery. Since most pre-mRNAs contain multiple introns,

different mRNAs can be produced from the same gene by different combinations of 5’ and 3’ splice

sites. The possibility of joining exons in varied combinations provides a novel means of controlling

gene expression by generating multiple mRNAs, and therefore multiple proteins, from the same pre-

mRNA. This process, known as alternative splicing, occurs frequently in genes of complex eukaryotes.

For example, it is estimated that about 50% of human genes produce transcripts that are alternatively

spliced, considerably increasing the diversity of proteins that can be encoded by the estimated 20,000-


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25,000 genes in mammalian genomes. Because patterns of alternative splicing can vary in different

tissues and in response to extracellular signals, alternative splicing provides an important mechanism

for tissue-specific and developmental regulation of gene expression.

One well-studied example of tissue-specific alternative splicing relates to sex determination in

Drosophila, where alternative splicing of the same pre-mRNA determines whether a fly is male or

female. Alternative splicing of the pre-mRNA of a gene called transformer is controlled by a protein

(SXL) that is only expressed in female flies. The transformer-derived pre-mRNA has three exons, but

different second exons are incorporated into the exon as a result of using alternate 3’ splice sites in the

two different sexes. In males, exon 1 is joined to the most upstream of these 3’ splice sites, which is

selected by the binding of the U2AF splicing factor. In females, the SXL protein binds to this 3’ splice

site, blocking the binding of U2AF. Consequently, the upstream 3’ splice site is skipped in females, and

exon 1 is instead joined to an alternate 3’ splice site that is further downstream. The exon 2 sequences

included in the male transformer mRNA contain a translation termination codon, so no protein is

produced. This termination codon is not included in the female mRNA, so female flies express a

functional transformer protein, which acts as a key regulator of sex determination.

The alternative splicing of transformer illustrates the action of a repressor (the SXL protein) that

functions by blocking the binding of a splicing factor (U2AF). Similarly, a large group of proteins

regulate alternative splicing by binding to silencer sequences in pre-mRNAs. In other cases, alternative

splicing is controlled by activators that recruit splicing factors to splice sites that would otherwise not

be recognized. The best-studied splicing activators are members of the SR protein family, which bind

to specific splicing enhancer sequences. Multiple mechanisms can thus regulate alternative splicing,

and variations in alternative splicing make a major contribution to the diversity of protein expression

during development and differentiation. One example is found in the mammalian ear, which contains

hair cells that respond to sounds of different frequencies. The responsiveness of hair cells to specific

frequencies is thought to be mediated in part by the alternative splicing of a gene encoding a channel

protein.

I. RNA EDITING

RNA editing refers to RNA processing events other than splicing that alter the protein-coding

sequences of some mRNAs. This unexpected form of RNA processing was first discovered in

mitochondrial mRNAs of trypanosomes in which U residues are added or deleted at multiple sites

along the pre-mRNA in order to generate the mRNA. More recently, editing has been described in

mitochondrial mRNA of other organisms, chloroplast mRNAs of higher plants, and nuclear mRNAs

of some mammalian genes.


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Editing in mammalian nuclear mRNAs, as well as in mitochondrial and chloroplast RNAs of

higher plants, involves single base changes as a result of base modification reactions, similar to those

involved in tRNA processing. In mammalian cells, RNA editing reactions include the deamination of

cytosine to uridine and of adenosine to inosine. One of the best-studied examples is the editing of the

mRNA for apolipoprotein B, which transports lipids in the blood. In this case, tissue-specific RNA

editing results in two different forms of apolipoprotein B. In humans, Apo-B100 is synthesized in the

liver by translation of the unedited mRNA. However, a shorter protein (Apo-B-48) is synthesized in

the intestine as a result of translation of an edited mRNA in which a C has been changed to a U by

deamination. This alteration changes the codon for glutamine (CAA) in the unedited mRNA to the

termination codon (UAA) in the edited mRNA, resulting in synthesis of the shorter Apo-B protein.

Tissue-specific editing of Apo-B mRNA thus results in the expression of structurally and functionally

different proteins in the liver and intestine. The full-length Apo-B100 produced by the liver transports

lipids in the circulation; Apo-B48 functions in the absorption of dietary lipids by the intestine. This

example is shown in Figure 36.21.

Figure 36.21: The effect of C-U RNA editing on the human ApoB gene


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RNA editing by the deamination of adenosine to inosine is the most common form of nuclear RNA

editing in mammals. This form of editing plays an important role in the nervous system, where A-to-I

editing results in single amino acid changes in ion channels and receptors on the surface of neurons.

For example, the mRNAs encoding receptors for the neurotransmitter serotonin can be edited at up to

five sites, potentially yielding 24 different versions of the receptor with different signaling activities.

J. CANCER

Cancer arises from a breakdown of the regulatory mechanisms that govern normal cell behaviors. As

discussed in preceding chapters, the proliferation, differentiation, and survival of individual cells in

multicellular organisms are carefully regulated to meet the needs of the organism as a whole. This

regulation is lost in cancer cells, which grow and divide in an uncontrolled manner either locally or

spread elsewhere throughout the body. This growth can seriously interfere with normal tissue and

organs.

Because cancer results from defects in fundamental cell regulatory mechanisms, it is a disease

that ultimately demands study at the molecular level. Indeed, understanding cancer has been a

paramount objective of molecular and cellular biologists for years. Importantly, studies of cancer cells

have also illuminated the mechanisms that regulate normal cell behavior. In fact, many of the proteins

that play a key role in cell signaling, regulation of the cell cycle, and control of programmed cell death

were first identified because abnormalities in their activities lead to the uncontrolled proliferation of

cancer cells, contributing to our understanding of normal cell regulation.

I. Tumor Viruses

Members of several families of animal viruses, called tumor viruses, are capable of directly causing

cancer in either experimental animals or humans. The viruses that cause human cancer include the

hepatitis B and C viruses, which cause liver cancer, papillomaviruses, which cause cervical and other

anogenital cancers, Epstein-Barr virus, which causes Burkitt’s lymphoma and nasopharyngeal

carcinoma, Kaposi’s sarcoma-associated herpesvirus, which causes Kaposi’s sarcoma, and human T-

cell lymphotropic virus, which causes adult T-cell leukemia. In addition, HIV is indirectly responsible

for the cancers that develop in AIDS patients as a result of immunodeficiency.

Tumor viruses, one of the earliest subjects of cancer research, came to serve as models for

cellular and molecular studies of cell transformation, the process by which normal cells are converted

to tumor cells. Their small genomes have made tumor viruses readily amenable to molecular analysis,


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leading to the identification of viral genes responsible for cancer induction and paving the way to our

current understanding of cancer at the molecular level.

Studies of tumor viruses demonstrated that specific genes, now known as oncogenes, are

capable of inducing cell transformation; these findings provided the first insights into the molecular

basis of cancer. However, more than 80% of human cancers are not induced by viruses and

apparently arise from other causes, such as radiation and chemical carcinogens. Therefore, in terms of

our overall understanding of cancer, it has been critically important that early studies of viral

oncogenes also led to the identification of cellular oncogenes.

II. Retroviral Oncogenes

Viral oncogenes were first defined in Rous sarcoma virus (RSV), a virus which transforms chicken

embryo fibroblasts in culture and induces large sarcomas within 1 to 2 weeks after inoculation into

chickens. In contrast, the closely related avian leucosis virus (ALV) replicates in the same cells as RSV

without inducing transformation. This difference in transforming potential suggested the possibility

that RSV contains specific genetic information responsible for transformation of infected cells. A direct

comparison of the genomes of RSV and ALV was consistent with this hypothesis: the genomic RNA

of RSV is about 10 kb, whereas that of ALV is smaller, about 8.5 kb.

In the early 1970s, a pair of researchers isolated deletion mutants and temperature-sensitive

mutants of RSV that were unable to induce transformation. Importantly, these mutants still replicated

normally in infected cells, indicating that RSV contains genetic information that is required for

transformation but not for virus replication. Further analysis demonstrated that both the deletion and

the temperature-sensitive RSV mutants define a single gene responsible for the ability of RSV to

induce tumors in birds and transform fibroblasts in culture. Because RSV causes sarcomas, its

oncogene is called src. The src gene is an addition to the genome of RSV; it is not present in ALV. It

encodes a 60-kd protein that was the first protein-tyrosine kinase to be identified.

More than 40 different highly oncogenic retroviruses have been isolated from a variety of

animals, including chickens, turkeys, mice, rats, cats, and monkeys. All of these viruses, like RSV,

contain either one or two oncogenes that are not required for virus replication but are responsible for

cell transformation. In some cases, different viruses contain the same oncogenes, but more than two

dozen distinct oncogenes have been identified among this group of viruses. Like src, many of these

genes, such as ras and raf, encode proteins that are now recognized as key components of signaling

pathways that stimulate cell proliferation.

An unexpected feature of retroviral oncogenes is their lack of involvement in viral replication.

Since most viruses are streamlined to replicate as efficiently as possible – this is, after all, their singular

evolutionary task – the existence of viral oncogenes that are not an integral part of the virus life cycle


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appears paradoxical. Scientists were thus led to question where the retroviral oncogenes had

originated and how they had come to be incorporated into viral genomes, a line of investigation that

ultimately led to the identification of cellular oncogenes in human cancers.

Scientists hypothesized that normal cells contain genes that are closely related to retroviral

oncogenes. The normal-cell genes from which the retroviral oncogenes originated are called proto-

oncogenes. These are important cell regulatory genes, in many cases encoding proteins that function

in the signal transduction pathways controlling normal cell proliferation (e.g., src, raf, and ras). The

oncogenes are abnormally expressed or mutated forms of the corresponding proto-oncogenes. As a

consequence of such alterations, the oncogenes induce abnormal cell proliferation and tumor

development.

An oncogene incorporated into a retroviral genome differs in several respects from the

corresponding proto-oncogene. First, the viral oncogene is transcribed under the control of viral

promoter and enhancer sequences, rather than being under the control of normal transcriptional

regulatory sequences. Consequently, oncogenes are usually expressed at a much higher level than the

proto-oncogenes, and are sometimes also expressed in inappropriate cell types. In some cases, such

abnormalities of gene expression are sufficient to convert a normally functioning proto-oncogene into

an oncogene that drives cell transformation.

In addition to such alterations in gene expression, oncogenes frequently encode proteins that

differ in structure and function from those encoded by their normal homologs. Many oncogenes, such

as raf, are expressed as fusion proteins with viral sequences at the amino terminus. Recombination

events leading to the generation of such fusion proteins often occur during and after the capture of

proto-oncogenes by retro-viruses, generating oncogene proteins that function in an unregulated

manner. For example, the viral raf oncogene encodes a fusion protein in which amino-terminal

sequences of the normal Raf protein have been deleted, as shown in Figure 36.22.


736

Figure 36.22: The Raf proto-oncogene protein consists of an amino-terminal

regulatory domain and a carboxy-terminal protein kinase domain. In the viral Raf

oncogene protein, the regulatory domain has been deleted and replaced by partially

deleted viral Gag sequences (Δ Gag). As a result, the Raf kinase domain is

constitutively active, causing cell transformation.

These amino-terminal sequences are critical to the regulation of the normal Raf protein kinase

activity, and their deletion results in unregulated constitutive activity of the oncogene-encoded Raf

protein. This unregulated Raf activity drives cell proliferation, resulting in transformation.

Many other oncogenes differ from the corresponding proto-oncogenes by point mutations,

resulting in single amino acid substitutions in the oncogene products. In some cases, such amino acid

substitutions (like the deletions already discussed) lead to unregulated activity of the oncogene proteins.

An important example of such point mutations is provided by the ras family of oncogenes, the role of

which are discussed in the next section.

III. Oncogenes in Human Cancer

Understanding the origin of retroviral oncogenes raised the question as to whether non-virus-induced

tumors contain cellular oncogenes that are generated from proto-oncogenes by mutations or by DNA

rearrangements during tumor development.

Some of the oncogenes identified in human tumors are cellular homologs of oncogenes that

were previously characterized in retroviruses, whereas others are new oncogenes first discovered in


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human cancers. The first human oncogene identified in gene transfer assays was identified as the

human homolog of the rasH oncogene of Harvey sarcoma virus. Three closely related members of the

ras gene family (rasH, rasK, and rasN) are the oncogenes most commonly encountered in human

tumors. These genes are involved in approximately 20% of all human malignancies, including more

than half of colon cancers and a quarter of lung carcinomas.

The ras oncogenes are not present in normal cells; rather, they are generated in tumor cells as

a consequence of mutations that occur during tumor development. The ras oncogenes differ from their

proto-oncogenes by point mutations resulting in single amino acid substitutions at critical positions. In

animal models, it has been shown that mutations that covert ras proto-oncogenes to oncogenes are

caused by chemical carcinogens, providing a direct link between the mutagenic action of carcinogens

and cell transformation.

The ras genes encode guanine nucleotide-binding proteins that function in transduction of

mitogenic signals form a variety of growth factor receptors. The activity of the Ras proteins is

controlled by GTP or GDP binding, such that they alternate between active (GTP-bound) and

inactive (GDP-bound) states. The mutations characteristic of ras oncogenes have the effect of

maintaining the Ras proteins constitutively in the GTP-bound conformation. In large part, this effect

is a result of nullifying the response of oncogenic Ras proteins to GAP (GTPase-activating protein),

which stimulates hydrolysis of bound GTP by normal RAS. Because of the resulting decrease in their

intracellular GTPase activity, the oncogenic Ras proteins remain in the active GTP-bound state and

drive unregulated cell proliferation.

Point mutations are only one of the ways in which proto-oncogenes are converted to

oncogenes in human tumors. Many cancer cells display abnormalities in chromosome structure,

including translocations, duplications, and deletions. The gene rearrangements resulting from

chromosome translocations frequently lead to the generation of oncogenes. In some cases, analysis of

these rearrangements has implicated already-known oncogenes in tumor development. In other cases,

novel oncogenes have been discovered by molecular cloning and analysis of rearranged DNA

sequences.

The first characterized example of oncogene activation by chromosome translocation was the

involvement of the c-myc oncogene in human Burkitt’s lymphoma, a malignancy of antibody-

producing B lymphocytes. The tumor is caused by chromosome translocations involving the genes that

encode immunoglobulins. For example, virtually all Burkitt’s lymphomas have translocations of a

fragment of chromosome 8 to one of the immunoglobulin gene loci which residue on chromosome 2 (κ

light chain), 14 (heavy chain), and 22 (λ light chain). One such translocation is shown in Figure 36.23.


738

Figure 36.23: The cy-myc proto-oncogene is translocated from chromosome 8 to

the immunoglobulin heavy-chain locus (IgH) on chromosome 14 in Burkitt’s

lymphomas, resulting in abnormal c-myc expression.

The fact that the immunoglobulin genes are actively expressed in these tumors suggested that the

translocation activates proto-oncogenes from chromosome 8 by inserting them into the

immunoglobulin loci. This possibility was investigated by analysis of tumor DNAs with probes for

known oncogenes, leading to the finding that the c-myc proto-oncogene was the chromosome 8

translocation break point in Burkitt’s lymphomas. These translocations inserted c-myc into an

immunoglobulin locus, where it was expressed in an unregulated manner. Such uncontrolled

expression of the c-myc gene, which encodes a transcription factor normally induced only in response

to growth factor stimulation, is sufficient to drive cell proliferation and contribute to tumor

development.

Translocations of other proto-oncogenes frequently result in rearrangements of coding

sequences, leading to the formation of abnormal gene products. The prototype for this process is

translocation of the abl proto-oncogene from chromosome 9 to chromosome 22 in chronic myeloid

leukemia (CML). This translocation leads to fusion of abl with its translocation partner, a gene called

bcr, on chromosome 22. The resulting product is the Bcr/Abl fusion protein in which the normal

amino terminus of the Abl proto-oncogene protein has been replaced by Bcr amino acid sequences.


739

The fusion of Bcr sequences results in unregulated activity of the Abl protein-tyrosine kinase, leading

to cell transformation. This translocation event is shown in Figure 36.24.

Figure 36.24: The abl oncogene is translocated from chromosome 9 to chromosome 22,

forming the Philadelphia chromosome in chronic myeloid leukemia (CML). The abl

proto-oncogene, which contains two alternative first exons (1A and 1B), is joined in the

middle of the bcr gene on chromosome 22. Exon 1B is deleted as a result of the

translocation. Transcription of the fused gene initiates at the bcr promoter and continues

through abl. Splicing then generates a fused Bcr/Abl mRNA in which abl exon 1A

sequences are joined to abl exon 2. The Bcr/Abl mRNA is translated to yield a

recombinant Bcr/Abl fusion protein.


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IV. Gene Amplification

A distinct mechanism by which oncogenes are activated in human tumors is gene amplification, which

results in elevated gene expression. DNA amplification is common in tumor cells, and amplification of

oncogenes may play a role in the progression of many tumors to more rapid growth and increasing

malignancy. Indeed, novel oncogenes have been identified by molecular cloning and characterization

of DNA sequences that are amplified in tumors.

A prominent example of oncogene amplification is the involvement of the N-myc gene, which

is related to c-myc, in neuroblastoma, a childhood tumor of neuronal cells. Amplified copies of N-myc

are frequently present in rapidly growing, aggressive tumors, indicating that N-myc amplification is

associated with the progression of neuroblastoma to increasing malignancy. Amplification of another

oncogene, erbB-2, which encodes a receptor protein-tyrosine kinase, is similarly related to progression

of breast and ovarian carcinomas.

I. Functions of Oncogene Products

The viral and cellular oncogenes encompass a large group (numbering more than 100 in total) that

can contribute to the abnormal behavior of malignant cells. As already noted, many of the proteins

encoded by proto-oncogenes regulate normal cell proliferation; in these cases, the elevated expression

or activity of the corresponding oncogene proteins drives the uncontrolled proliferation of cancer cells.

Other oncogene products contribute to the behavior of cancer cells as well, such as failure to undergo

programmed cell death or defective differentiation.

The function of oncogene proteins in regulation of cell proliferation is illustrated by their

activities in growth factor-stimulated pathways of signal transduction, such as the activation of ERK

signaling downstream of receptor protein-tyrosine kinases. The oncogene proteins within this pathway

include polypeptide growth factors, growth factor receptors, intracellular signaling proteins,

transcription factors, and the cell cycle regulatory cyclin D1.

The action of growth factors as oncogene proteins results from their abnormal expression,

leading to a situation where a tumor cell produces a growth factor to which it also responds. The result

is autocrine stimulation of the growth factor-producing cell, which drives abnormal cell proliferation

and contributes to the development of a wide variety of human tumors.

A large group of oncogenes encode growth factor receptors, most of which are protein-

tyrosine kinases. These receptors can be converted to oncogene proteins by alterations of their amino-

terminal domains, which would normally bind extracellular growth factors. For example, the receptor

for platelet-derived growth factor (PDGF) is converted to an oncogene in some human leukemias by a

chromosome translocation in which the normal amino terminus of the PDGF receptor (PDGFR) is


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replaced by the amino terminal sequence of a transcription factor called Tel. Alternatively, genes that

encode receptors are induced in response to protein-tyrosine kinases and can be activated by gene

amplification or by point mutations that result in unregulated kinase activity. Other oncogenes

(including src and abl) encode nonreceptor protein-tyrosine kinases that are constitutively activated by

deletions or mutations of regulatory sequences.

The Ras proteins play a key role in mitogenic signaling by coupling growth factor receptors to

activation of the Raf protein-serine/threonine kinase, which initiates a protein kinase cascade leading

to activation of ERK MAP kinase. As discussed already, the mutations that convert ras proto-

oncogenes to oncogenes results in constitutive Ras activity, thus causing activation of the ERK and

PI3K/Akt pathways. The raf gene can similarly be converted to an oncogene by deletions that result in

loss of the amino-terminal regulatory domain of the Raf protein. The consequence of these deletions is

unregulated activity of the Raf protein kinase, which also leads to constitutive ERK activation.

Alternatively, raf proto-oncogenes can be converted to oncogenes by point mutations that result in

elevated Raf kinase activity. A schematic of this effector pathway, as well as its potential points of

dysregulation, is shown in Figure 36.25.

Figure 36.25: Canonical Raf effector pathway. Points of mutations implicated in the

development and progression of human cancers, involving Raf-MEK-ERK or PI3K-Akt

pathway dysregulation, are shown.


742

The ERK pathway ultimately leads to the phosphorylation of transcription factors and

alternations in gene expression. As might be expected, many oncogenes encode transcriptional

regulatory proteins that are normally induced in response to growth factor stimulation. For example,

transcription of the fos proto-oncogene is induced as a result of phosphorylation of Elk-1 by ERK. Fos

and the product of another proto-oncogene, Jun, are components of the AP-1 transcription factor,

which activates transcription of a number of target genes, including that which gives rise to cyclin D1,

in growth factor-stimulated cells. Constitutive activity of AP-1, resulting from unregulated expression

of either the Fos or Jun oncogene proteins, is sufficient to drive abnormal cell proliferation, leading to

cell transformation. The Myc proteins similarly function as transcription factors regulated by

mitogenic stimuli, and abnormal expression of myc oncogenes contributes to the development of a

variety of human tumors. Other transcription factors are activated as oncogenes by chromosome

translocations in human leukemias and lymphomas.

The signaling pathways activated by growth factor stimulation ultimately regulate

components of the cell cycle machinery that promote progression through the restriction point G1.

The D-type cyclins are induced in response to growth factor stimulation, at least in part via activation

of the AP-1 transcription factor. These proteins play a key role in coupling growth factor signaling to

cell cycle progression. Perhaps not surprisingly, the gene encoding cyclin D1 is a proto-oncogene,

which can be activated as the oncogene CCND1 by chromosome translocation or gene amplification.

These alterations lead to constitutive expression of cyclin D1, which then drives cell proliferation in

the absence of normal growth factor stimulation. The catalytic partner of cyclin D1, Cdk4, is also

activated as an oncogene, by point mutations in melanomas.

Components of other signaling pathways, including the G protein-coupled signaling

pathways, the NF-κB pathway, and the Hedgehog, Wnt, and Notch pathways, can also act as

oncogenes. For example, activating mutations frequently convert the downstream target of Wnt

signaling, β-catenin, to an oncogene (CTNNB1) in human colon cancers. These activating mutations

stabilize β-catenin, which then forms a complex with Tcf and stimulates transcription of target genes.

The targets of β-catenin/Tcf include the genes encoding c-Myc and cyclin D1, leading to unregulated

cell proliferation. Interestingly, Wnt signaling normally promotes the proliferation of stem cell and

their progeny during the continual epithelial cell renewal that occurs in the colon, indicating that

colon cancer results from abnormal activity of the same pathway that signals physiologically normal

proliferation of colonic epithelial cells.

Although many oncogenes stimulate cell proliferation, the oncogenic activity of some

transcription factors instead results from inhibition of cell differentiation. As noted elsewhere, thyroid

hormone and retinoic acid induce differentiation of a variety of cell types. These hormones diffuse

through the plasma membrane and bind to intracellular receptors that act as transcriptional regulatory

mechanisms. A mutated form of the retinoid acid receptor (PML/RARα) acts as an oncogene protein


743

in human acute promyelocytic leukemia. The mutated oncogene receptor appear to interfere with the

action of its normal homologs, thereby blocking cell differentiation and maintaining the leukemic cells

in an actively proliferating state. In the case of acute promyelocytic leukemia, high doses of retinoic

acid can overcome the effects of the PML/RARα protein oncogene protein and induce differentiation

of the leukemic cells. This biological observation has a direct clinical correlation: patients with acute

promyelocytic leukemia can be treated effectively by administration of retinoic acid, which induces

differentiation and blocks continued cell proliferation.

As already emphasized, the failure of cancer cells to undergo programmed cell death

(apoptosis) is a hallmark of cancerous cells. Several oncogenes code proteins that act to promote cell

survival, which, in most animal cells, is dependent on growth factor stimulation. Accordingly, those

oncogenes that encode growth factors, growth factor receptors, and signaling proteins such as Ras act

not only to promote cell proliferation, but also to prevent cell death. The PI3-kinase/Akt signaling

pathway plays an anti-apoptotic role in many growth factor-dependent cells, and the genes encoding

PI3-kinase and Akt act as oncogenes in both retroviruses and human tumors. The downstream targets

of PI3-kinase/Akt signaling include a proapoptotic member of the Bcl-2 family Bad, which is

inactivated as a result of phosphorylation of Akt, as well as the FOXO transcription factor, which

regulates expression of the proapoptotic Bcl-2 family member, Bim. In addition, it is worth mentioning

that Bcl-2 itself was first discovered as the product of an oncogene in human lymphomas. The bcl-2

oncogene is generated by a chromosome translocation that results in elevated expression of Bcl-2,

which blocks apoptosis and maintains cell survival under conditions that normally induce cell death in

the development of cancer.

II. Tumor Suppressor Genes

The activation of cellular oncogenes represents only of two distinct types of genetic alterations involved

in tumor development; the other is inactivation of tumor suppressor genes. Oncogenes drive abnormal

cell proliferation as a consequence of genetic alterations that either increase gene expression or lead to

uncontrolled activity of the oncogene-encoded proteins. Tumor suppressor genes represent the

opposite effect—they control growth, normally acting to inhibit cell proliferation and tumor

development. In many tumors, these genes are lost or inactivated, thereby removing negative

regulators of cell proliferation and contributing to the abnormal proliferation of tumor cells. The

functions of tumor suppressor proteins encoded by tumor suppressor genes can be broadly delineated

into one of several categories:


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♦ Repression of genes that are essential for the continuance of the cell cycle. If

these genes are not expressed, the cell cycle does not continue, effectively

bringing the cell cycle, and cell division, to a halt.

♦ Coupling the cell cycle to DNA damage. As long as there is damaged DNA in the

cell, it should not divide; the cell cycle should continue only if the detected

damage is repaired.

♦ If the damage cannot be repaired, the cell should initiate apoptosis (programmed

cell death) within the damaged cells.

♦ Some proteins involved in cell adhesion prevent tumor cells from dispersing,

block loss of contact inhibition, and inhibit metastasis. These proteins are known

as metastasis suppressors.

♦ DNA repair proteins are usually classified as tumor suppressors as well, as

mutations in their genes increase the risk of cancer. Examples of these mutations

include HNPCC, MEN1 and BRCA. Furthermore, increased mutation rate from

decreased DNA repair leads to increased inactivation of other tumor suppressors

and activation of oncogenes.

The tumor suppressor gene was identified by studies of retinoblastoma, a rare childhood

ophthalmic tumor. Provided that the disease is detected early, retinoblastoma can be successfully

treated and many patients survive to have families. For this reason, it was recognized that same cases

of retinoblastoma appear to be inherited. In these cases, approximately 50% of the children of an

affected parent develop retinoblastoma, consistent with Mendelian transmission of a single dominant

gene that confers susceptibility to tumor development. Although susceptibility to retinoblastoma is

transmitted as a dominant trait, inheritance of the susceptibility gene is not sufficient to transform a

normal retinal cell into a tumor cell.

Cell transformation associated with the development of retinoblastoma requires two

mutations, which are now known to correspond to the loss of both functional copies of the tumor

susceptibility gene (the Rb tumor suppressor gene) that would be present on homologous chromosomes

of a normal diploid cell. In inherited retinoblastoma, one defective copy of Rb is genetically

transmitted. The loss of this single Rb copy is not by itself sufficient to trigger tumor development, but

retinoblastoma almost always develops in these individuals as a result of a second mutation leading to

the loss of the remaining normal Rb allele. Noninherited retinoblastoma, in contrast, is rare, since its


745

development requires two independent somatic mutations to inactivate both normal copies of Rb in

the same cell.

The observation that alleles that code for Rb must be affected before an effect is manifested is

more generally referred to as the “two-hit hypothesis.” This is because if only one allele for a tumor

suppressor gene is damaged, the second can still produce the correct protein. In other words, mutant

tumor suppressors' alleles are usually recessive whereas mutant oncogene alleles are typically

dominant.

The two-hit hypothesis was first proposed by A.G. Knudson for cases of retinoblastoma, when

Knudson observed that the age of onset of retinoblastoma followed second order kinetics. This pattern

implied that two independent genetic events were necessary. Knudson recognized that this was

consistent with a recessive mutation involving a single gene, but requiring bi-allelic mutation.

Oncogene mutations, in contrast, generally involve a single allele because they are gain-of-function

mutations.

The functional nature of the Rb gene as a negative regulator of tumorigenesis is not isolated to

retinoblastoma; it is also involved in more common adult tumors. In particular, studies of the cloned

gene have established that Rb is lost or inactivated in many bladder, breast, and lung carcinomas. The

significance of the Rb tumor gene thus extends beyond retinoblastoma, and mutations of the Rb gene

contribute to a substantial fraction of human cancers. The Rb protein is a key target for the oncogene

proteins of several DNA tumor viruses, including SV40, adenoviruses, and human papillomaviruses,

which bind to Rb and inhibit its activity. Transformations by these viruses thus result, at least in part,

from inactivation of Rb at the protein level, rather than from mutational inactivation of the Rb gene.

Characterization of Rb as a tumor suppressor gene served as the conceptual catalyst for

research that identified many additional tumor suppressor genes that contribute to the development of

a host of human malignancies. Some of these genes were identified as the cause of rare inherited

cancers, playing a role similar to that of Rb in hereditary retinoblastoma. Other tumor suppressor

genes have been identified as genes that are frequently deleted or mutated in common noninherited

cancers of adults, such as colon carcinoma. In either case, evidence strongly supports the proposition

that tumor suppressor genes are involved in the development of both inherited and noninherited forms

of cancer. In fact, mutations of some tumor suppressor genes appear to be the most common

molecular alterations leading to human tumor development.

The second tumor suppressor gene to have been identified is p53, which is frequently

inactivated in a wide variety of human cancers, including leukemias, lymphomas, sarcomas, brain

tumors, and carcinomas of many tissues, including breast, colon, and lung. Certain mutations in the

p53 gene product represent an exception to the “two-hit” rule for tumor suppressors. p53 mutations

can function as a ‘dominant negative,’ meaning that a mutated p53 protein can prevent the function of

normal protein from the un-mutated allele.


746

In total, mutations of p53 play at least some role in more than half of all cancers, making it the most

common target of genetic alterations in human malignancies. It is also of interest that inherited

mutations of p53 are responsible for genetic transmission of a rare hereditary autosomal dominant

cancer syndrome, Li-Fraumeni syndrome, in which affected individuals develop any of several

different types of cancers. In addition, the p53 protein (like RB) is a target for the oncogene proteins of

SV40, adenoviruses, and human papillomaviruses. In one such example, human papillomavirus

(HPV), encodes a protein, E6, which binds to the p53 protein and inactivates it. This mechanism, in

synergy with the inactivation of the cell cycle regulator Rb by the HPV protein E7, allows for repeated

cell division manifested clinically as warts. Certain HPV types, in particular types 16 and 18, can also

lead to progression from a benign wart to low or high-grade cervical dysplasia, which are reversible

forms of precancerous lesions. Persistent infection of the cervix can cause irreversible changes leading

to carcinoma in situ and eventually invasive cervical cancer. This results from the effects of HPV genes,

particularly those encoding E6 and E7—two viral oncoproteins that are preferentially retained and

expressed in cervical cancers by integration of the viral DNA into the host genome.

The p53 protein is continually produced and degraded in cells of healthy people. The

degradation of the p53 protein is associated with binding of MDM2. In a negative feedback loop,

MDM2 itself is induced by the p53 protein. Mutant p53 proteins often fail to induce MDM2, causing

p53 to accumulate at very high levels. Moreover, the mutant p53 protein itself can inhibit normal p53

protein levels. In some cases, single missense mutations in p53 have been shown to disrupt p53 stability

and function. The function of MDM2 as part of the p53 pathway in a normal cell is shown in Figure

36.26.


747

Figure 36.26: In a normal cell, p53 is inactivated by its negative regulator, mdm2. Upon

DNA damage or other stresses, various pathways will lead to the dissociation of the p53-

mdm2 complex. Once activated, free p53 will induce either cell cycle arrest to repair the

cell or apoptosis to discard the damaged cell. The mechanism by which one of these

pathways is selected by p53 is an area of ongoing research.

Like p53, the INK4 and PTEN tumor suppressor genes are very frequently mutated in several

common cancers, including lung and prostate cancers and melanoma. Other tumor suppressor genes

(including APC, TβRII, Smad2, and Smad4) are frequently inactivated in colon cancers. In addition to

being involved in non-inherited cases of this common adult cancer, inherited mutations of the APC

gene are responsible for a rare hereditary form of colon cancer, called familial adenomatous polyposis

(FAP). Individuals with this condition develop hundreds of benign colon adenomas (polyps), some of

which inevitably progress to malignancy. Inherited mutations of two other tumor suppressor genes,

BRCA1 and BRCA2, are responsible for hereditary cases of breast cancer, which account for about 5%


748

of total breast cancer incidence. Additional tumor suppressor genes have been implicated in the

development of brain tumors, pancreatic cancer, and basal skin carcinomas, as well as several rare

inherited cancer syndromes, such as Wilms’ tumor.

III. Products of Tumor Suppressor Genes

In contrast to proto-oncogene and oncogene proteins, the proteins encoded by most tumor suppressor

genes inhibit cell proliferation or survival. Inactivation of tumor suppressor genes therefore leads to

tumor development by eliminating negative regulatory proteins. In many cases, tumor suppressor

proteins inhibit the same regulatory pathways that are stimulated by the products of oncogenes.

The protein encoded by PTEN tumor suppressor gene is an interesting example of

antagonism between oncogene and tumor suppressor gene products. The PTEN protein is a lipid

phosphatase that dephosphorylates the 3 position of phosphatidylinositides, such as

phosphatidylinositol 3,4,5-bisphosphate (PIP3). By dephosphorylating PIP3, PTEN antagonizes the

activities of PI3-kinase and Akt, both of which can act as oncogenes by promoting cell survival.

Conversely, inactivation or less of the PTEN tumor suppressor protein can contribute to tumor

development as a result of increased levels of PIP3 and Akt, and inhibition of programmed cell death.

Proteins encoded by both oncogenes and tumor suppressor genes also function in the

Hedgehog signaling pathway. The receptor Smoothened is an oncogene in basal cell carcinomas,

whereas Patched (the negative regulator of Smoothened) is a tumor suppressor gene.

Several tumor suppressor genes encode transcriptional regulatory proteins. A good example is

the product of WT1, which is frequently inactivated in Wilms’ tumor (a childhood renal tumor). The

WT1 protein is a repressor that appears to suppress transcription of a number of growth-factor

inducible genes. One of the targets of WT1 is thought to be the gene that encodes an insulin-like

growth factor, which is overexpressed in Wilms’ tumor and may contribute to tumorigenesis by acting

as an autocrine growth factor. Inactivation of WT1 may thus lead to abnormal growth factor

expression, which in turn drives cell proliferation. Two other tumor suppressor genes, Smad2 and

Smad4, encode transcription factors that are activated by TGF-β signaling and lead to inhibition of cell

proliferation. Consistent with the activity of TGF-β in inhibiting cell proliferation, the TGF-β receptor

is also encoded by a tumor suppressor gene (TβRII).

The products of the Rb and INK4 tumor suppressor genes regulate cell cycle progression at the

same point as that affected by cyclin D1 and Cdk4, both of which can act as oncogenes. Rb inhibits

progression through the restriction point in G1 by repressing transcription of a number of genes

involved in cell cycle progression and DNA synthesis. In normal cells, passage through the restriction

point is regulated by Cdk4,6/cyclin D complexes, which phosphorylate and inactivate Rb. Mutational

inactivation of Rb in tumors thus removes a key negative regulator of cell cycle progression. The INK4


749

tumor suppressor gene, which encodes the Cdk inhibitor p16, also regulates passage through the

restriction point; in normal cells, p16 inhibits Cdk4,6/cyclin D activity. Inactivation of INK4 therefore

leads to elevated activity of Cdk4,6/cyclin D complexes, resulting in uncontrolled phosphorylation of

Rb.

The p53 gene product regulates both cell cycle progression and apoptosis. DNA damage leads

to rapid induction of p53, which activates transcription of both proapoptotic and cell cycle inhibitory

genes. The effect of p53 on apoptosis are mediated in part by activating the transcription of

proapoptotic members of the Bcl-2 family—PUMA and Noxa—that induce programmed cell death.

Unrepaired DNA damage normally induces apoptosis of mammalian cells; this response is presumably

advantageous to the organism because it eliminates cells carry potentially deleterious mutations,

including cells that may develop into cancerous cells. Cells lacking p53 fail to undergo apoptosis in

response to agents that damage DNA, including radiation and many of the drugs used in

chemotherapy treatment. This failure to undergo apoptosis in response to DNA damage contributes to

the resistance of many tumor cells to chemotherapeutic agents. In addition, loss of p53 appears to

interfere with apoptosis induced by other stimuli, such as growth factor deprivation and oxygen

deprivation. These effects of p53 on cell survival are thought to account for the high frequency of p53

mutations in human cancers.

In addition to inducing apoptosis, p53 bocks cell cycle progression in response to DNA

damage by inducing the Cdk inhibitor p21. The p21 protein blocks cell cycle progression by acting as

a general inhibitor of Cdk/cyclin complexes, and the resulting cell cycle arrest presumably allows time

for damaged DNA to be repaired before it is replicated. Loss of p53 prevents this damage-induced cell

cycle arrest, leading to increasing mutation frequencies and a general instability of the cellular

genome. Such genetic instability is a common property of cancer cells, and it may further contribute to

alterations in oncogenes and tumor suppressor genes during tumor progression. These regulatory

mechanisms, as well as other pathways previously discussed, are shown in Figure 36.27.


750

Figure 36.27: RAS induces the transcriptional upregulation of growth factors and

interferes with transforming growth factor-β (TGFβ) signaling through inhibition of

TGFβ receptor expression or downstream signaling by downregulating the expression of

SMAD3, as well as the nuclear accumulation of SMAD2 and SMAD3. RAS also

upregulates the levels of cyclin D1 and suppresses the cyclin-dependent kinase inhibitor

(CDKI) p27. The newly synthesized cyclin D1 associates with and activates the cyclin-

dependent kinases CDK4 and CDK6, leading to the phosphorylation of RB and the

subsequent dissolution of the RB–E2F transcription factor complexes. Once released,

E2F transcription factors transactivate several genes that are required for cell cycle

progression, including cyclin E (CCNE) and cyclin A (CCNA) that induce transition

through the G1/S checkpoint (not shown). Hyperproliferative cues from activation of the

RAS oncogene can result in replicative stress leading to DNA damage. In response to

DNA damage, cells can activate the DNA damage checkpoints to transiently arrest and

restore the integrity of the genome, enter a state of irreversible arrest (senescence) or

undergo apoptosis. Inaccurate repair of DNA damage can lead to mutations and

chromosome aberrations, thereby contributing to tumorigenesis. The asterisk represents

the mutational activation of RAS; P represents phosphorylation.

Although their function remains to be fully understood, the products of the BRCA1 and

BRCA2 genes (which are responsible for some inherited breast and ovarian cancers) also appear to be

involved in checkpoint control of cell cycle progression and repair of double-stranded breaks in DNA.

BRCA1 and BRCA2 thus function as stability genes, acting to maintain the integrity of the genome.

Mutations in genes of this type lead to the development of cancer not as a result of direct effects on cell

proliferation or survival, but because their inactivation leads to a high frequency of mutations in


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oncogenes or tumor suppressor genes. Other stability genes whose loss contributes to the development

of human cancers include the ATM gene, which acts as a DNA damage checkpoint, the mismatch

repair genes that are defective in some inherited colorectal cancers, and the nucleotide excision repair

genes that are mutated in the dermatological condition known as xeroderma pigmentosum.


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Chapter 36 Problems


In investigating the avian retrovirus RSV, researchers noted that infection of chicken fibroblast cells by

RSV led to neoplastic transformation of the infected host cell. Their previous research indicated that a

single viral gene, src, was responsible. Because many highly oncogenic retroviruses were isolated from

the tumors of infected animals, they hypothesized that retroviral oncogenes are derived from related

genes of host cells. Consistent with this suggestion, normal cells of several species were found to

contain retrovirus-related DNA sequences that could be detected by nucleic acid hybridization, but

that could not alone lead to cell transformation. However, it was unclear whether these sequences

were related to the retroviral oncogenes or to the genes required for virus replication.

To address this question, the researchers isolated transformation-defective mutants of RSV that

sustained a 1.5 kb deletion corresponding to most or all of the src gene. They synthesized a radioactive

DNA probe composed of short single-stranded DNA (ssDNA) fragments complementary to the entire

genomic RNA of normal RSV. This probe was then hybridized to an excess of RNA isolated from

transformation-defective mutants. Fragments of cDNA that were complementary to the viral

replication genes hybridized to the transformation-defective RSV RNA. In contrast, cDNA fragments

that were complementary to src were unable to hybridize and remained single-stranded.

The radioactive src cDNA was then used as a hybridization probe to attempt to detect related DNA

sequences in normal avian fibroblast cells not infected by RSV. The extent of hybridization of src

cDNA to normal chicken, quail and duck DNA is shown in Figure 1. When introduced into non-avian

fibroblast cells, little cDNA hybridization occurred, but transformation was still noted in a large

percentage of observed cells.


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Figure 1: Hybridization of src-specific cDNA to normal chicken, quail, and duck DNA. (Note: final

cDNA hybridization > 50% was considered reflective of significant sequence homology.)

1. Given the size of the deletion observed in the transformation-defective mutants isolated by the

researchers, what is the best prediction regarding a property of the unhybridized region of the src-

specific probe?

A. It is homologous with approximately 1.5 kb of RSV RNA taken from a normal RSV virus.

B. It is homologous with approximately 1.5 kb of RSV RNA in the transformation-defective RSV

mutant.

C. It is homologous with approximately 1.5 kb of cDNA derived from the RNA taken from a

transformation-defective RSV mutant.

D. Its nucleotide composition is identical to that of a 1.5 kb sequence of RNA taken from a normal

RSV virus.

2. Which result most supports the hypothesis that retroviral oncogenes are derived from related genes

present in host cells?

A. cDNA fragments that were complementary to the viral replication genes hybridized to the

transformation-defective RSV RNA.


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B. cDNA fragments that were complementary to src were unable to hybridize to any segment of RSV

RNA.

C. src cDNA probes hybridized extensively to normal avian DNA.

D. cDNA formed RNA-DNA duplexes when hybridized to transformation-defective RSV RNA.

3. Researchers postulated that the structural similarity between the src gene and homologous

sequences found in normal avian fibroblasts, known as c-src, exists because of the incorporation of host

cell DNA into an ancestral RSV virus. If the researchers are correct, which of the following is most

likely to be true of c-src?

A. Host cell DNA transferred to the RSV ancestor possessed the independent ability to transform

avian fibroblast cells prior to the transfer.

B. RSV causes host cell transformation only when c-src is present.

C. Mutations leading to the oncogenic potential of the src gene occurred in RSV.

D. The structural similarity observed is due to transfection of the host cell by RSV with mutated host

cell DNA.

4. Following mutation, the gene to which the cDNA probe hybridized in avian cells promotes

angiogenesis, cell proliferation and cell migration. Before modification, such a gene is best described as

a(n):

A. oncogene.

B. tumor suppressor gene.

C. reporter gene.

D. proto-oncogene.


5. Which of the following is NOT true regarding DNA footprinting?

A. It is used to identify protein binding sites on DNA.

B. The DNA sequence is labelled at both ends of the tested strands.

C. The site at which the protein binds will be protected from digestion by DNase.

D. Fragments are subject to electrophoresis following digestion.


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6. Which of the following statements correctly describes the functionally and structurally different

forms of apolipoprotein B synthesized in the human liver and intestine?

A. Apo-B100 is synthesized in the liver by translation of the unedited mRNA transcript.

B. Apo-B48 is synthesized in the intestine by translation of an edited mRNA transcript in which the

editing reaction has eliminated a stop codon.

C. Apo-B100 is synthesized in the liver by translation of an edited mRNA transcript in which the

editing reaction has generated a stop codon.

D. Apo-B48 is synthesized in the intestine by translation of the unedited mRNA transcript.

7. Which of the following is true of X chromosome inactivation in human females?

I. One of the two X chromosomes is inactivated early in female development.

II. Xist is produced from a gene on one of the two X chromosomes.

III. Xist recruits proteins that induce condensation of chromatin.

A. I only

B. II only

C. II and III only

D. I, II and III

8. What is the principle function of splicing factors that are not components of snRNPs?

A. They introduce double-stranded breaks into unedited mRNA transcripts.

B. They mediate the process by which mRNAs that lack open reading frames are degraded.

C. They direct snRNPs to the correct splice site.

D. They recognize GC-rich inverted repeat segments of mRNA molecules.


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9. Which of the following regulatory elements is responsible for the division of individual domains of

chromatin into fixed spatial regions and prevents the action of trans-acting elements in one domain

from interacting with regulatory elements in an adjacent domain?

A. Repressor

B. Promoter

C. Enhancer

D. Insulator

10. Can a proto-oncogene be converted to an oncogene without a change or mutation in its coding

sequence?

A. No, a proto-oncogene can only be converted to an oncogene via an activating sequence change.

B. Yes, a proto-oncogene can be activated by a translocation that puts it under the control of a

mutated, inactive promoter sequence.

C. Yes, a proto-oncogene can be activated by a mutation that silences a tumor suppressor gene.

D. Yes, a proto-oncogene may be expressed in abnormal cell types.

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1. A.

According to the passage, the transformation-defective mutants of RSV sustained a 1.5 kb deletion

corresponding to most or all of the src gene. The researchers synthesized a radiolabeled DNA probe

composed of short ssDNA fragments complementary to the genome of normal RSV, and hybridized it

to RNA isolated from the transformation-defective mutant lacking a 1.5 kb segment corresponding to

the RSV src gene. For this reason, it is expected that the segment of the probe which does not

hybridize with RSV RNA-lacking src must be complementary to the region of normal RSV RNA-

containing src and correspond to the 1.5 kb deletion in the transformation-defective mutant. This is

consistent with choice A, and eliminates choices B and C. In the case of choice D, the unhybridized

region will be derived from and complementary to a 1.5 kb sequence of RNA in the normal RSV

virus, but, as it is composed of DNA, its nucleotide composition will not be identical. Choice D is thus

incorrect.

2. C.

The observation stated in choice C is reflected in the text of the passage and in Figure 1. The src

cDNA probes derived from RSV hybridized extensively to normal avian DNA, indicating strong

sequence similarity between the oncogenic src gene of RSV and sequences in normal avian DNA. This

further supports the hypothesis that src in RSV was derived from a similar gene in normal avian DNA.

This conclusion is most similar to the idea referred to in the question. Choices A, B and D are

incorrect, as they only reflect the sequence similarities or differences, respectively, between normal and

transformation-defective RSV. They show no relationship between RSV sequences and

complementary sequences found in normal avian DNA.

3. C.

It is possible that at one point an ancestral virus mistakenly incorporated the c-src gene of its cellular

host. Eventually this normal gene mutated into an abnormally functioning oncogene within RSV.

Once the oncogenic mutated virus (known as v-src) is transfected back into a chicken, it can lead to

cancer. Choice C indicates a step in this sequence of events. The passage indicates that sequences

complementary to src in normal avian cells do not alone lead to cell transformation. This contradicts

choice A. Choice B is also unlikely to be true. The passage states that when introduced into non-avian

fibroblast cells, little cDNA hybridization occurred, but transformation was still noted in a large

percentage of observed cells. This indicates that transformation by the RSV virus did not depend on

the presence of c-src or another structurally similar sequence. Finally, choice D is also false. The


significant hybridization between the cDNA probe and normal avian fibroblast cells indicates that

structural similarity between src and c-src exists in cells not infected by, and therefore not previously

transfected with, RSV-related genetic material.

4. D.

Promotion of angiogenesis, cell migration and cellular proliferation are consistent with events seen in

cell transformation and cell reproduction. An oncogene is a gene that normally directs cell growth

which may be associated with cancer. A proto-oncogene is a normal gene that can become an

oncogene due to mutations or increased expression. Proto-oncogenes code for proteins that help to

regulate cell growth and differentiation and as such are often involved in signal transduction and

execution of mitogenic signals, usually through their protein products. Upon mutagenic activation, a

proto-oncogene (or its gene product) becomes a tumor-inducing agent, an oncogene. Prior to

modification, the gene was not affecting cell growth processes but after mutation, it did, making choice

A incorrect and choice D, a proto-oncogene, the correct designation. A tumor suppressor gene is a

gene that protects a cell from transformation. The mutation of tumor suppressor genes, however,

would not directly lead to the promotion of transformative characteristics in cells, including those

mentioned in the question, making choice B incorrect. Choice C is incorrect; a reporter is a gene that

a scientist will attach to a regulatory sequence of another gene of interest in the course of an

experiment. Certain genes are chosen as reporters because the characteristics they confer on

organisms expressing them (via protein production) are easily identified and measured, or because they

are selectable markers. Reporter genes are used as an indication of whether a certain gene has been

taken up by, or expressed in, the cell or organism studied.

5. B.

As discussed in the chapter, during DNA footprinting the DNA sequence is radiolabelled at one end

only. The statements in choices A, C and D are all true of the normal footprinting process, and are

incorrect.

6. A.

Apo-B100 is synthesized in the liver by translation of the unedited mRNA transcript. Apo-B48 is

synthesized in the intestine by translation of an edited mRNA transcript in which the editing reaction

has generated a premature stop codon. Thus choice A is correct and choices B, C and D are incorrect.

7. D.

Roman numerals I, II and III are all true. One of the two X chromosomes is inactivated early in

female development (or in that of XXY males). An RNA called Xist is produced by the Xist gene on

Chapter 36: Eukaryotic Chromosome Organization and Gene Expression Control

759

one of the two X chromosomes, and binds to most of the genes located on the chromosome. Xist RNA

recruits proteins that induce chromatin condensation and conversion of most of the inactive X

material to heterochromatin.

8. C.

Choice C is correct. Splicing factors that are not components of snRNPs direct snRNPs to the correct

splice sites by binding to specific sequences in the pre-mRNA.

9. D.

Insulators are barrier regulatory elements that divide chromosomes into individual domains of

chromatin structure that can be either chromatin or heterochromatin, but that cannot spread beyond

the insulator. They can also prevent an enhancer in one domain from acting on a promoter in an

adjacent domain. This is choice D. Choice A is incorrect. A repressor is a DNA- or RNA-binding

protein that inhibits the expression of one or more genes by binding to the operator or associated

silencers. Choices B and C are also incorrect. A promoter is a region of DNA that initiates

transcription of a particular gene. Promoters are located near the transcription start sites of genes, on

the same strand and upstream on the DNA (towards the 5' region of the sense strand). An enhancer is

a short region of DNA that can be bound with proteins (activators) to activate transcription of a gene

or genes. These proteins are usually referred to as transcription factors.

10. D.

If expressed in an abnormal cell type, a proto-oncogene or its protein product(s) may function

oncogenically in the cell, even if it would not do so when expressed in a proper cell type. This is choice

D. Choice A is false for the reason given in support of choice D. Choices B and C are also false; a

proto-oncogene can be activated without a change or mutation in its coding sequence by gene

amplification, or if a translocation event puts it under the control of an activator. However, the loss of

a functional tumor suppressor gene alone does not confer oncogenic character on a proto-oncogene.

Some further mutation converting the proto-oncogene to an oncogene is ordinarily still required.


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Chapter 37 Recombinant DNA and Biotechnology

A. INTRODUCTION

Classical experiments in molecular biology were strikingly successful in developing our

fundamental concepts of the nature and expression of genes. Since these studies were based primarily

on genetic analysis, their success depended largely on the choice of simple, rapidly replicating

organisms; bacteria and viruses served as frequent model systems. It was not clear, however, how these

fundamental principles could be extended to the complexities inherent in eukaryotic cells, since the

genomes of most eukaryotes are thousands of times larger than those of bacteria. These obstacles were

overcome by the development of recombinant DNA technology, which provided scientists with a

means of isolating, sequencing and manipulating genes derived from any type of cell. The application

of recombinant DNA technology has thus enabled detailed molecular studies of the structure and

function of eukaryotic genes and genomes, thereby revolutionizing our understanding of molecular

and cell biology.

Recombinant DNA technology, also called gene cloning or molecular cloning, is a general

term that encompasses a number of experimental protocols leading to the transfer of DNA from one

organism to another. There is no single method that can be used to satisfy this objective. However, a

recombinant DNA experiment often follows a similar sequence:

♦ The DNA, whether cloned, inserted, targeted, or foreign, from a donor organism

is extracted, enzymatically cleaved, and joined to another DNA entity (a cloning

vector). This forms a new, recombinant DNA molecule, often called a DNA

construct.

♦ The DNA construct is transferred into and maintained within a host cell. The

introduction of DNA into a bacterial host cell is called transformation.

♦ Those host cells that take up the DNA construct (transformed cells) are identified

and selected by separation or isolation from untransformed cells.

♦ If required, a DNA construct can be created so that the protein product encoded

by the cloned DNA sequence is produced in the host cell.

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801

B. RESTRICTION ENZYMES

The first step in the development of recombinant DNA technology was the characterization of

restriction endonucleases, which are enzymes that cleave DNA at specific sequences. These enzymes

were identified in bacteria, where they apparently provide a defense against the entry of foreign DNA

(e.g., from a virus) into the cell. Bacteria have a variety of endonucleases that cleave DNA at hundreds

of distinct recognition sties, each of which consists of a specific sequence of four to eight base pairs.

Many of these nucleotide sequences are palindromic, meaning the base sequence reads the same

backward and forward. An example of such a sequence is shown in Figure 37.1.

Figure 37.1: The sequence of nucleotides in a palindromic recognition site is the same in

the forward and reverse strands when both are read in the same 5’ to 3’ or 3’ to 5’

orientation.

Two types of palindromic sequences exist in DNA. A mirror-like palindrome is similar to that

which could be found in ordinary text, in which a sequence is the same when read in the forward and

backward directions on a single strand of DNA, as in GTAATG. The inverted repeat palindrome is

also a sequence that reads the same in both directions, but the forward and backward sequences are

found in complementary DNA strands (i.e., of double-stranded DNA), as in GTATAC (GTATAC

being complementary to CATATG). The sequence shown in Figure 37.1 is an inverted repeat

palindrome. Inverted repeat palindromes are more common and have greater biological importance

than mirror-like palindromes.

Restriction enzymes also differ in terms of the structure of the ends of the double-stranded

DNA (dsDNA) fragments which they produce. Certain restriction enzymes form so-called “sticky

ends”; others form “blunt ends.” The simplest end of a double stranded DNA molecule is called a

blunt end. In a blunt-ended molecule, both strands terminate in a base pair.

Non-blunt ends are created by various overhangs. An overhang is a stretch of unpaired

nucleotides in the end of a DNA molecule. These unpaired nucleotides can be in either strand,

creating either 3' or 5' overhangs. In most cases, these overhangs are palindromic. An example is given

in Figure 37.2.


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Figure 37.2: Single nucleotide overhang

The simplest case of an overhang is a single nucleotide. This is most often adenosine, which is

established as a 3' overhang by some DNA polymerases. Most commonly, this is used in cloning PCR

products created by such an enzyme. The product is joined with a linear DNA molecule with 3'

thymine overhangs. Since adenine and thymine form a base pair, this facilitates the joining of the two

molecules by a ligase, yielding a circular molecule.

Longer overhangs are called cohesive ends or sticky ends. They are most often created by

restriction endonucleases when they cut DNA. Often, they cut the two DNA strands four base pairs

from each other, creating a four-base 5' overhang in one molecule and a complementary 5' overhang

in the other. These ends are called cohesive since they are easily rejoined by a ligase. Also, since

different restriction endonucleases usually create different overhangs, it is possible to cut a piece of

DNA with two different enzymes before ligating it to a DNA molecule with ends formed by the same

enzymes. Since the overhangs have to be complementary in order for the ligase to function properly,

the two molecules can only join in one orientation. Sticky-end restriction digests are shown in Figure

37.3.

Figure 37.3: Sticky ends formed by restriction digestion can base pair in

complementary overhang regions.

Naturally occurring restriction endonucleases are categorized into four groups (types I, II, III,

and IV) based on their composition and enzyme cofactor requirements, the nature of their target

sequence, and the position of their DNA cleavage sites relative to the target sequence. The

differentiating characteristics of these endonuclease are detailed below.


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♦ Type I enzymes cleave at sites remote from a recognition site. They require both

ATP and S-adenosyl-L-methionine to function. They are multifunctional proteins

with both restriction and methylase activities.

♦ Type II enzymes cleave within or at short specific distances from recognition

sites; most require magnesium. They are single function restriction enzymes

lacking methylase activity.

♦ Type III enzymes cleave at sites a short distance from a recognition site. They

require ATP, but do not hydrolyze it. S-adenosyl-L-methionine stimulates the

reactions which they catalyze, but is not required for the enzymes to function. The

enzymes exist as part of a complex with a modification methylase that modifies

existing methylated residues in protein.

♦ Type IV enzymes target modified DNA, specifically methylated,

hydroxymethylated and glucosyl-hydroxymethylated DNA.

Since restriction endonucleases digest DNA at specific sequences, they can be used to cleave a

DNA molecule at unique sites, forming fragments of variable lengths. These digested fragments can be

separated according to size by gel electrophoresis, as shown in Figure 37.4.


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Figure 37.4: Restriction enzyme digestion results in cleavage at specific sequence sites

(shown as grey arrows) in DNA. These fragments are then separated by electrophoresis in

an agarose gel. The DNA fragments migrate toward the positive electrode (anode), with

smaller fragments moving more rapidly through the gel. Following electrophoresis, the

DNA is stained with a fluorescent dye and photographed. The sizes of DNA fragments

are indicated.

The location of cleavage sites for multiple different restriction endonucleases can be used to

generate detailed restriction maps of DNA molecules, such as viral genomes. In addition, individual

DNA fragments, produced by restriction endonuclease digestion, can be isolated following

electrophoresis for further study, including determination of their DNA sequence. The DNA of many

viruses has been characterized by this approach.

Restriction endonuclease digestion alone, however, does not provide sufficient resolution for

the analysis of larger DNA molecules, such as cellular genomes. A restriction endonuclease with a six-


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base-pair recognition site (such as the enzyme EcoRI) cleaves DNA with a statistical frequency of once

every 4096 base pairs. Digestion of the human genome, which is more than three million base pairs

long, would yield more than 500,000 EcoRI fragments. Because it is impossible to isolate single

restriction fragments from such a large pool of digests, restriction endonuclease digestion alone does

yield a homogenous source of DNA suitable for further analysis. Such quantities can, however, be

obtained through molecular cloning.

C. MOLECULAR CLONING

As introduced earlier, the basic strategy in molecular cloning is to insert a DNA fragment of

interest (e.g. a segment of human DNA) into a DNA molecule that serves as a vector. Such a vector

must be capable of independent replication within a host cell. The result is a recombinant molecule or

molecular clone, composed of the DNA insert linked to vector DNA sequences. Large quantities of the

inserted DNA can be obtained if the recombinant molecule is allowed to replicate in an appropriate

host. For example, fragments of human DNA can be cloned in plasmid vectors. This is shown in

Figure 37.5. Plasmids are small circular DNA molecules that can replicate independently in bacteria;

in other words, they can do so without being associated with chromosomal DNA. Recombinant

plasmids carrying human DNA inserts can be introduced into E. coli, where they replicate along with

the bacteria to yield millions of copies of plasmid DNA. The DNA of these plasmids can then be

isolated, generating large quantities of recombinant molecules containing a single fragment of human

DNA. The fragment can then easily isolated from the rest of the vector DNA by restriction

endonuclease digestion and gel electrophoresis, allowing a pure fragment of human DNA to be

analyzed and further manipulated.

The DNA fragments used to create recombinant molecules are usually generated by

digestion with restriction endonucleases. Many of these enzymes cleave their recognition sequences at

staggered sites, leaving overhanging or cohesive single-stranded tails that can associate with each other

by complementary base pairing. The association between such paired complementary ends can be

established permanently by treatment with DNA ligase. Thus, two different fragments of DNA (e.g. a

human DNA insert and a plasmid DNA vector) prepared by digestion with the same restriction

endonuclease can be readily joined to create a recombinant DNA molecule.

The fragments of DNA that can be cloned are not limited to those that terminate in

restriction endonuclease cleavage sites. Synthetic DNA “linkers” containing desired restriction

endonuclease sites can be added to the ends of a DNA fragment, allowing virtually any fragment of

DNA to be ligated to a vector and isolated as a molecular clone.


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Figure 37.5: Generation of a recombinant DNA molecule. 1. Small, circular DNA

molecules called plasmids are removed from bacterial cells. These plasmids serve as

vectors carrying genes of interest. This plasmid includes antibiotic resistance genes, a

reporter gene responsible for coloration, LacZ, and within the LacZ gene, a multiple

cloning site (also known as a polylinker) containing various restriction sites. 2. Foreign

DNA containing the gene of interest is extracted from the cell. 3. A restriction enzyme


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recognizes its specific restriction site – a short sequence 4-8 base pairs long. 4. The

foreign DNA is cleaved, producing fragments with sticky ends. The restriction enzyme

cuts and opens the circular plasmids. The same enzyme cuts the gene of interest from its

DNA molecule. 5. The sticky ends anneal by forming weak hydrogen bonds. Adding

DNA ligase reattaches the DNA backbones and results in the formation of a recombinant

plasmid. Other plasmids reseal, and are unchanged. 6. Plasmids and bacteria and bacteria

are mixed. Many of the bacteria do not take any plasmids into their cells, many take

plasmids without the foreign DNA in them, and a few will take up the recombinant

plasmid via transformation. 7. Plasmids with an uninterrupted LacZ gene are blue. In the

recombinant plasmids, the inserted gene interrupts the LacZ gene, and the bacteria remain

their original color. Bacteria which do not take up any plasmids also remain uncolored.

Antibiotics are then added and because the plasmid contains the genes for antibiotic

resistance, only bacteria which have incorporated the plasmid survive the antibiotic. The

bacteria can now be sorted according to color, isolating the bacteria which took up the

plasmid containing the gene of interest and uncolored bacteria are allowed to reproduce.

I. cDNA

Cloning is not limited to DNA sequences; RNA sequences can be cloned as well. The first

step is to synthesize a DNA copy of the RNA using the enzyme reverse transcriptase. The DNA

product is called cDNA because it is complementary to the template RNA. cDNA can then be ligated

to vector DNA in the manner previously discussed. Since eukaryotic genes are usually interrupted by

noncoding sequences, which are removed from mRNA by splicing, the ability to clone cDNA as well

as genomic DNA has been critical for understanding gene structure and function. Additionally, cDNA

cloning allows the mRNA corresponding to a single gene to be isolated as a molecular clone.


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II. Vectors in Recombinant DNA

Depending on the size of the insert DNA and the purpose of the experiment, many different

types of cloning vectors can be used for the generation of recombinant molecules. We will review some

of those here. Other vectors developed for the expression of cloned DNAs and the introduction of

recombinant molecules into eukaryotic cells will be discussed in subsequent sections.

Plasmids are commonly used for cloning genomic or cDNA inserts of up to a few

thousand base pairs. Plasmids usually consist of 2 to 4 kb of DNA, including an origin of replication,

which is the DNA sequence that signals the host cell DNA polymerase to replicate the DNA molecule.

In addition, plasmid vectors carry genes that confer antibiotic resistance, so bacteria carrying the

plasmids can be selected. For example, Figure 37.6 illustrates the isolation of human cDNA clones in a

plasmid vector.


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Figure 37.6: The vector is a small circular molecule that contains an origin of replication

(ori), a gene conferring resistance to ampicillin (Ampr), and a restriction site, which can

be used to insert foreign DNA. Insert DNA (in this case, a human cDNA fragment) is

ligated to the vector, and the recombinant plasmids are used to transform the vector. The

bacteria are plated on medium containing ampicillin, so that only the bacteria that are

ampicillin-resistant because they carry plasmid DNAs are able to form colonies (not

pictured).

A pool of cDNA fragments can be ligated to restriction endonuclease-digested plasmid DNA.

The resulting recombinant DNA molecules are then used to transform E. coli. Antibiotic-resistant

colonies which contain plasmid DNA are selected by exposure to the antibiotic for which the

transformed bacteria possess resistance. Since each recombinant plasmid yields a single antibiotic-

resistant colony, the bacteria present in any given colony will contain a unique cDNA insert. Plasmid-

containing bacteria can then be grown in large quantities and their DNA extracted. The small circular

plasmid DNA molecules, of which there are often hundreds of copies per cell, can be separated from

the bacterial chromosomal DNA; the result is purified plasmid DNA that is suitable for analysis of the

cloned insert.

Bacteriophage λ vectors are also used for the isolation of either genomic or cDNA

clones from eukaryotic cells, and will accommodate larger fragments of insert DNA than plasmids. In λ

cloning vectors, sequences of the bacteriophage genome that are dispensable for virus replication have

been removed and replaced with unique restriction sites for insertion of cloned DNA. These

recombinant molecules can be introduced into E. coli, where they replicate to yield millions of progeny

phages containing a single DNA insert. The DNA of these phages can then be isolated, yielding large

quantities of recombinant molecules containing a single fragment of cloned DNA. The DNA inserts

can be as large as 15 kb and still typically yield a recombinant genome that can be packaged into

bacteriophage λ particles.

For many studies involving analysis of genomic DNA, it is desirable to clone larger

fragments than are accommodated by plasmid or λ vectors. There are five major types of vectors that

are used for this purpose. Cosmid vectors (plasmid vectors that contain cos sites, sites which

circularizes the DNA in the host cytoplasm) accommodate inserts of approximately 45 kb. These

vectors contain bacteriophage λ sequences that allow efficient packaging of the cloned DNA into phage

particles. In addition, cosmids contain origins of replications and the genes for antibiotic resistance

that are characteristic of plasmids, so they are able to replicate as plasmids in bacterial cells. Two other

types of vectors are derived from bacteriophage P1, rather than from bacteriophage λ. Bacteriophage

P1 vectors, which will accommodate DNA fragments of 70-100 kb, contain sequences that allow

recombinant molecules to be packaged in vitro into P1 phage particles and then to be replicated as


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plasmids in E. coli. P1 artificial chromosome (PAC) vectors also contain sequences of bacteriophage P1,

but are introduced directly as plasmids into E. coli. They will accommodate larger inserts of up to 150

kb. Bacterial artificial chromosome (BAC) vectors are derived from a naturally occurring plasmid of E.

coli—the F factor. The replication origin and other F factor sequences allow BACs to replicate as

stable plasmids carrying inserts of 120-300 kb. Even larger fragments of DNA (250-400 kb) can be

cloned in yeast artificial chromosome (YAC) vectors. These vectors contain yeast origins of replication

as well as other sequences, including centromere and telomeres that allow replication as linear

chromosome-like molecules in yeast cells.

D. DNA SEQUENCING

Molecular cloning allows the isolation of individual fragments of DNA in quantities suitable

for detailed characterization, including the determination of nucleotide sequence. Indeed, determining

the nucleotide sequence of many genes has elucidated not only the structure of their products, but also

the properties of DNA sequences that regulate gene expression. Furthermore, the coding sequences of

novel genes are frequently related to those of previously studied genes, and the functions of newly

isolated genes can often be correctly deduced on the basis of such sequence similarity.

DNA sequencing is usually performed with automated systems that are both rapid

and accurate, so determining the sequence of several kilobases of DNA is a straightforward task. Thus,

it is easier to clone and sequence DNA than it is to determine the amino acid sequence of a protein.

Since the nucleotide sequence of a gene can be readily translated into the amino acid sequence of its

encoded protein, the easiest way of determining protein sequence is the sequencing of a cloned gene or

cDNA.

I. Sanger Chain-Termination Method

The most common method of DNA sequencing is based on premature termination of DNA

synthesis resulting from the inclusion of chain-terminating dideoxynucleotides, which do not contain

the 3’ hydroxyl group, in DNA polymerase reactions. DNA synthesis is initiated at a unique site on the

cloned DNA from a synthetic primer. The DNA synthesis reaction includes each of four

dideoxynucleotides (A, C, G, and T) in addition to their normal counterparts. Each of the four

dideoxynucleotides is labeled with a different fluorescent dye, so their incorporation into DNA can be

monitored. Incorporation of these dideoxynucleotides stops further DNA synthesis because no 3’

hydroxyl group is available as a site for the addition of the next nucleotide. Thus, a series of labeled

DNA molecules is generated, each terminating at the base represented by a specific dideoxynucleotide.

Those fragments of DNA are then separated according to size by gel electrophoresis. As the newly


811

synthesized DNA strands are electrophoresed through the gel, they pass through a laser that excites

the fluorescent labels. The resulting emitted light is then detected by a photomultiplier, and a

computer collects and analyzes the resultant data. The size of each fragment is determined by its

terminal dideoxynucleotide, marked by a specific color fluorescence, so the DNA sequence can be

read from the order of fluorescent-labeled fragments as they migrate through the gel. High-throughput

automated DNA sequencing of this type has enabled large-scale analysis required for determination of

the sequences of completed genomes, include that of humans. This process is summarized in Figure

37.7.

Figure 37.7: Dideoxynucleotides, which lack OH groups at the 3’ as well as the 2’

position of deoxyribose, are used to terminate DNA synthesis at specific bases. These

molecules are incorporated normally into growing DNA strands. Because they lack a 3’

OH, however, the next nucleotide cannot be added, so synthesis of that DNA strand

terminates. DNA synthesis is initiated at a specific site with a primer. The reaction

contains the four dideoxynucleotides. When the dideoxynucleotides is incorporated,

DNA synthesis stops, so the reaction yields a series of products extending from the

primer to the base substituted by a fluorescent dideoxynucleotide. These products are

then separated by gel electrophoresis. As the DNA strands migrate through the gel, they


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pass through a laser beam that excites the fluorescent labels on the dideoxynucleotides.

The emitted light is detected by a photomultiplier, which is connected to a computer that

collects and analyzes the data to determine the sequence of DNA.

E. EXPRESSING CLONED GENES

In addition to enabling determination of the nucleotide sequences of genes – and hence the

amino acid sequences of their protein products – molecular cloning has provided new approaches to

obtaining large amounts of proteins for structural and functional characterization. Many proteins of

interest are present at only low levels in eukaryotic cells and therefore cannot be purified in significant

amounts by conventional biochemical techniques. Given a cloned gene, however, this problem can be

rectified by the engineering of vectors that lead to high levels of gene expression in either bacteria or

eukaryotic cells.

To express a eukaryotic gene in E. coli, the cDNA of interest is cloned into a plasmid

or phage vector (called an expression vector) that contains sequences that drive the expression of the

inserted gene in bacterial cells. Inserted genes often can be expressed at levels high enough that the

protein encoded by the cloned gene corresponds to as much as 10% of the total bacterial protein

complement. Purifying the protein encoded by the cloned gene in quantities suitable for detailed

biochemical or structural studies is a straightforward matter.

It is frequently useful to express high levels of a cloned gene in eukaryotic cells rather

than in bacteria. This mode of expression may be important, for example, to ensure that

posttranslational modifications of the protein, such as additions of carbohydrates or lipids, occur

normally. This protein expression in eukaryotic cells can be achieved, as in E. coli, by insertion of the

cloned gene into a (usually virally-derived) vector that directs high-level gene expression. One system

frequently used for protein expression in eukaryotic cells is infection of insect cell baculovirus vectors,

wherein exceedingly high levels of expression of genes inserted in place of a viral structural protein

occurs. Alternatively, high levels of protein expression can be achieved using appropriate vectors in

mammalian cells.

Expression of cloned genes in yeast is particularly useful because simple methods of

yeast genetics can be employed to identify proteins that interact with one another. In this type of

analysis, known as the yeast two-hybrid system, two different cDNAs (for example, from human cells)

are joined to two distinct domains of a protein that stimulates expression of a target gene in yeast.

Figure 37.8 illustrates a yeast two-hybrid system.


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Figure 37.8: A yeast-two hybrid system. cDNAs of two human proteins are cloned and

fused with two distinct domains of a yeast protein that stimulates transcription of a target

gene. The two recombinant cDNAs are introduced into a yeast cell. Domain 1 binds DNA

sequences at a site upstream of the target gene, and domain 2 stimulates target gene

transcription. The interaction between the two human proteins can thus be detected by

expression of the target gene in transformed yeast.

Yeast are then transformed with hybrid cDNA clones to test for interactions between the two

proteins. If the human proteins interact with each other, they will bring the two domains of the yeast

protein together, resulting in stimulation of target expression in the transformed yeast. Expression of

the target gene can be easily detected by the growth of yeast in a specific medium or by the production

of an enzyme that produces a blue yeast colony, so the yeast two-hybrid system provides a

straightforward method to evaluate protein-protein interactions. Indeed, high-throughput yeast two-

hybrid screens have been used to construct large-scale interaction maps of thousands of proteins in

eukaryotic cells.

F. DETECTION OF NUCLEIC ACIDS

The advent of molecular cloning has enabled the isolation and characterization of individual

genes from eukaryotic cells. Understanding the roles of genes within cells, however, requires analysis of

intracellular organization and expression of individual genes and their encoded proteins. In this

section, the basic procedures used for detection of specific nucleic acids will be discussed. These

approaches are important for a wide variety of studies, including the mapping of genes to

chromosomes and the analysis of gene expression.

I. Amplification of DNA by the Polymerase Chain Reaction


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Molecular cloning allows individual DNA fragments to be propagated in bacteria and isolated

in large amounts. An alternative method for isolating large amounts of a single DNA molecule is the

polymerase chain reaction (PCR). Provided that some sequence of the DNA molecule of interest is

known, PCR can achieve a striking amplification of DNA content via reactions carried out entirely in

vitro. Essentially, DNA polymerase is used for repeated replications of a defined segment of DNA. The

number of DNA molecules increases exponentially, doubling with each round of replication, so a

substantial quantity of DNA can be obtained from a relatively small initial sample of template copies.

For example, a single DNA molecule amplified through 30 cycles of replication would theoretically

yield 230, or more than a billion, progeny molecules. Single DNA molecules can thus be amplified to

yield readily detectable quantities of DNA that can be isolated by molecular cloning or further

analyzed directly by restriction endonuclease digestion or nucleotide sequencing. The general

procedure for PCR amplification is shown in Figure 37.9.

Figure 37.9: The target region of DNA to be amplified is flanked by two strands used to

prime DNA synthesis. In the first step of each cycle, the starting dsDNA is separated and

then cooled to allow the primers, usually oligonucleotides 15-20 bases long, to bind to

each strand of ssDNA. Taq polymerase is used to synthesize new DNA strands from the

primers, resulting in the formation of two new DNA molecules. The process can be

repeated for multiple cycles, each resulting in a twofold amplification of DNA. 1.

Denaturation at 94-96°C. 2. Annealing at approximately 68°C. 3. Elongation at

approximately 72°C.


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The starting material in PCR amplification of DNA can be either a cloned fragment

of DNA or a mixture of DNA molecules – for example, total DNA from human cells. A specific region

of DNA can be amplified from such a mixture, provided that the nucleotide sequence surrounding the

region is known so that primers can be designed to initiate DNA synthesis at the desired point. Such

primers are usually chemically synthesized oligonucleotides containing 15-20 bases of DNA. Two

primers are used to initiate DNA synthesis in opposite directions from complementary DNA strands.

The reaction is started by heating the template DNA to a high temperature, typically 95°C, to

separate the two strands. The temperature is then lowered to allow the primers to pair with their

complementary sequences on the template strands. DNA polymerase then uses the primers to

synthesize a new strand complementary to each template. Thus, through a single cycle of

amplification, two new DNA molecules are synthesized from one template molecule. This process can

be repeated multiple times, with a twofold increase in the number of DNA molecules following each

round of replication.

The multiple cycles of heating and cooling involved in PCR are performed by

programmable heating blocks called thermocyclers. The DNA polymerases used in these reactions are

heat-stable enzymes from bacteria such as Thermus aquaticus, which reside in hot springs where

temperatures can exceed 75° C. (DNA polymerase derived from Thermus aquaticus is called Taq

polymerase.) Because these polymerases remain stable even at high temperatures, they are used to

separate the strands of DNA in double-stranded DNA, so PCR amplification can be performed

rapidly and automatically. RNA sequences can also be amplified by this method if reverse

transcriptase is used to synthesize a cDNA copy prior to PCR amplification.

If the sequence of a target gene is known sufficiently well, a primer for it can be

specified. Given this, PCR amplification provides a powerful tool for detecting small amounts of

specific DNA or RNA molecules in a complex mixture of other molecules. In such a situation, the only

DNA molecules that will be amplified by PCR are those containing sequences complementary to the

primers used in the reaction. Therefore, PCR can selectively amplify a specific template from

heterogeneous mixtures, such as total cell DNA or RNA. This extraordinary sensitivity has made PCR

an important method for a variety of applications, including analysis of gene expression in cells where

target DNA is available in only small quantities. The DNA segments amplified by PCR can also be

directly sequenced or ligated to vectors and propagated as molecular clones. PCR thus allows the

amplification and cloning of any segment of DNA for which primers can be designed. Since the

complete genome sequences of many organisms are now known, PCR can be used to amplify and

clone a wide array of desired DNA fragments.

II. Nucleic Acid Hybridization


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Another tool in the repertoire of molecular biologists takes advantage of the specific base

pairing between complementary strands of DNA or RNA. At high temperatures (90-100°C), the

complementary strands of DNA denature, yielding single-stranded DNA (ssDNA). If such ssDNA

molecules are then incubated under appropriate conditions, at temperatures close to 65°C, they will

re-nature and reform dsDNA as dictated by the pattern of complementary base pairing; this process is

called nucleic acid hybridization. Nucleic acid hybrids can be formed between two strands of DNA,

two strands of RNA, or one strand of DNA and one of RNA.

As discussed above, hybridization between the primers and the template DNA provides the

specificity of PCR amplification. In addition, a variety of other methods use nucleic acid hybridization

as a means for detecting DNA or RNA sequences that are complementary to any isolated nucleic acid,

such as a cloned DNA sequence. The cloned DNA is labeled with either radioactive nucleotides or

with modified nucleotides that can be detected by fluorescence or chemiluminescence. This labeled

DNA is then used as a probe for hybridization to complementary DNA or RNA sequences, which are

detected by virtue of the radioactivity, fluorescence, or luminescence of the resulting double-stranded

hybrids.

III. Southern Blotting

Southern blotting is widely used for the detection of specific genes in cellular DNA. The DNA

to be analyzed is digested with a restriction endonuclease, and the digested DNA fragments are

separated by gel electrophoresis. The gel is then overlaid with nitrocellulose filter paper or a nylon

membrane to which the DNA fragments are blotted (transferred) to yield a replica of the gel. The filter

is then incubated with a labeled probe, which hybridizes to the DNA fragments that contain the

complementary sequence, allowing visualization of these specific fragments of DNA. The steps of the

Southern blotting procedure are shown in Figure 37.10.


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Figure 37.10: Southern blotting. 1. DNA is digested by restriction endonuclease treatment

and the resultant restriction fragments of different sizes are separated by gel

electrophoresis. 2. The DNA is denatured and transferred to a filter by passage of a salt

solution through the gel. 3. The filter is hybridized with a labeled probe, which binds to

complementary DNA sequences in buffer solution. 4. The probe bound to the filter is

detected by exposure to film, which reveals the DNA fragments to which the probe

hybridized.

The capillary blotting system shown in step 2 from Figure 37.10 is shown in more detail

in Figure 37.11.


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Figure 37.11: Capillary blotting system used for the transfer of DNA from an

electrophoresis gel to a blotting membrane.

IV. Northern Blotting

As you may have guessed from its name, Northern blotting is a variation of the Southern

blotting technique. It is used for the detection of RNA, rather than DNA. In this method, total cellular

RNAs are extracted and fractionated according to size by gel electrophoresis. As in Southern blotting,

the RNAs are transferred to a filter and detected by hybridization with a cloned probe. Northern

blotting is frequently used in studies of gene expression – for example, to determine whether specific

mRNAs are present in different types of cells. The general procedure for Northern blotting is shown in

Figure 37.12.

Figure 37.12: Northern blotting. 1. RNA is extracted from sample. 2. RNA is fractionated

by size via gel electrophoresis. 3. RNA is transferred to filter. 4. RNA is fixed to

membrane when exposed to UV radiation or heat. 5. Labeled probes are added. 6.

Labeled RNA is visualized on x-ray film.


819

Nucleic acid hybridization can also be used to identify molecular clones that contain specific

cellular DNA inserts. The first step in isolation of either genomic or cDNA clones is frequently the

preparation of recombinant DNA libraries, collections of clones that contain all the genomic or

mRNA sequences of a particular cell type. For example, a genomic library of human DNA might be

prepared by cloning random DNA fragments of about 15 kb in a λ vector. Since the human genome is

approximately 3 x 106 kb, the complete human genome would be represented in a collage of

approximately 500,000 such clones. Any gene for which a probe is available can then be isolated from

such a recombinant library.

The recombinant phages are plated on E. coli, and each phage replicates to produce a plaque

on the lawn of bacteria. The plaques are then blotted onto filter in a process similar to the transfer of

DNA from a gel to a filter during Southern blotting, and the filters are hybridized with a labeled probe

to identify the phage plaques that contain the gene of interest. A variety of probes can be used for such

experiments. For example, a cDNA clone can be used as a probe to isolate the corresponding genomic

clone, or a gene cloned from one species (e.g., mouse) can be used to isolate a related gene from a

different species (e.g., human). The appropriate plaque can then be isolated from the original plate in

order to propagate the recombinant phage that carries the desired DNA insert. Similar procedures can

be used to screen bacterial colonies carrying plasmid DNA clones, so specific clones can be isolate by

hybridization from either phage or plasmid libraries. Figure 37.13 shows a protocol for screening a

recombinant library by hybridization.


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Figure 37.13: Screening of recombinant DNA libraries by the colony hybridization

procedure. 1. Fragments of cell DNA are cloned in a bacteriophage λ vector and

packaged into phage particles, yielding recombinant phage carrying different cell inserts.

2. The phage are used to infect bacteria, forming plaques. 3. The culture is overlaid with

filter paper; some of the phages in each plaque are transferred to the filter. 4. The phage

DNA is then hybridized with a labeled probe to identify the phage plaque containing the

desired gene. The appropriate phage plaque can then be isolated from the original culture

plaque.

V. DNA Microarrays

Rather than analyzing one gene at a time, as in Southern or Northern blotting, hybridization

to DNA microarrays allows tens of thousands of genes to be analyzed simultaneously. As the complete

sequences of eukaryotic genomes have become available, hybridization of DNA microarrays has

enabled researchers to undertake global analyses of sequences present in either cellular DNA or RNA

samples. A DNA microarray consists of a glass slide or membrane filter on which oligonucleotides or

fragments of cDNA are printed by a robotic system in small spots at high density. Each spot on an


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array consists of a single oligonucleotide or cDNA. More than 10,000 unique DNA sequences can be

printed onto a typical glass microscope slide, so it is readily possible to produce DNA microarrays

containing sequences representing all of the genes in cellular genomes. One widespread application of

DNA microarrays is in the study of gene expression; for example, it can be used to compare the genes

expressed by two different cell types. In an experiment of this type, cDNA probes are synthesized from

the mRNAs expressed in each of the two cell types (e.g. cancer cells and normal cells). The two cDNAs

are labeled with different fluorescent dyes (typically red and green) and a mixture of the cDNAs is

hybridized to a DNA microarray in which 10,000 or more human genes are represented as single

spots. The array is then analyzed using a high-resolution laser scanner, and the relative extent of

transcription of each gene in the cancer cells compared to the normal cells is indicated by the ratio of

red to green fluorescence at the a given position on the array. This procedure is shown in Figure

37.13.

Figure 37.13: DNA microarrays. An example of comparative analysis of gene expression

in cancerous cells and normal cells is shown. mRNAs extracted from cancer cells and


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normal cells are used as templates for synthesis of cDNA probes labeled with different

fluorescent dyes. Here, cDNA derived from cancer cells carries a red fluorescent label

and cDNA from normal cells carries a green label. The two cDNA probes are mixed and

hybridized to a DNA microarray containing spots of oligonucleotides corresponding

collectively to 10,000 or more distinct human genes. The relative expression of each gene

in cancer cells compared to normal cells is indicated by the ratio of red to green

fluorescence at each position on the microarray.

V. In situ Hybridization

Nucleic acid hybridization can be used to detect homologous DNA or RNA sequences not

only in cell extracts, but also in chromosomes or intact cells; this procedure called in situ hybridization.

In this case, the hybridization of fluorescent probes to specific cells or subcellular structures is analyzed

by microscopic examination. For example, labeled probes can be hybridized to intact chromosomes in

order to identify the chromosomal regions that contain a gene of interest. In situ hybridization can also

be used to detect specific mRNAs in different types of cells within a tissue.

G. UNDERSTANDING GENE FUNCTION IN EUKARYOTES

The recombinant DNA techniques discussed in the preceding sections provide powerful

approaches to the isolation and detailed characterization of the genes of eukaryotic cells.

Understanding the function of those genes, however, requires analysis of the gene within cells or intact

organisms, not simply as a molecular clone in bacteria. In classical genetics, gene function has

generally been revealed by the altered phenotypes of mutant organisms. The advent of recombinant

DNA has added a new dimension to studies of this function. Namely, it has become possible to

investigate the function of a cloned gene directly by reintroducing the cloned DNA into eukaryotic

cells. In simpler eukaryotes, such as yeasts, this technique has made possible the isolation of molecular

clones corresponding to virtually any mutant gene. In addition, there are several methods by which

cloned genes can be introduced into cultured animal and plant cells, as well as intact organisms, for

functional analysis. These approaches can be coupled with the ability to introduce mutations in cloned

DNA in vitro, extending the power of recombinant DNA to allow functional studies of the genes of

more complex eukaryotes.

I. Genetic Analysis in Yeasts


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Yeasts are particularly advantageous for studies of eukaryotic molecular biology. The genome

of Saccharomyces cerevisiae, which consists of approximately 1.2 x 107 base pairs, is nearly 200 times

smaller than the human genome. Moreover, yeasts can easily be grown in culture, reproducing with a

division time of about 2 hours. Thus yeasts offer the same basic advantages – a small genome and

rapid reproduction – that are afforded by bacteria.

Furthermore, mutations in yeast can be identified as readily as in E. coli. For example,

yeast mutants that require a particular amino acid or other nutrient for growth can easily be isolated.

In addition, yeasts with defects in genes required for fundamental cell processes (in contrast to

metabolic defects) can be isolated as temperature-sensitive mutants. Such mutants encode proteins that

are functional at one temperature (the permissive temperature) but not another (the non-permissive

temperature); in contrast, normal proteins are functional at both. A yeast with a temperature-sensitive

mutation in an essential gene can be identified by its ability to grow only at the permissive

temperature. The ability to isolate such temperature-sensitive mutants has allowed the identification of

yeast genes controlling many fundamental cell processes, such as RNA synthesis and processing,

progression through the cell cycle, and transport of proteins between cellular compartments.

The relatively simple genetics of yeast also enables a gene corresponding to any yeast

mutation to be cloned on the basis of its functional activity. First, a genomic library of normal yeast

DNA is prepared in vectors that replicate as plasmids in yeasts as well as in E. coli. The small size of the

yeast genome means that a complete library consists of only a few thousand plasmids. A mixture of

such plasmids is then used to transform a temperature-sensitive yeast mutant, and transformants that

are able to grow at the non-permissive temperature are selected. Such transformations have acquired a

normal copy of the gene of interest on plasmid DNA, which can then be easily isolated from the

transformed yeast cells for further characterization.

Yeast genes encoding a wide variety of essential proteins have been identified in this

manner. In many cases, such genes isolated from yeasts have also been useful in identifying and

cloning related genes from mammalian cells. Thus, the simple genetics of yeast has not only provided

an important model for eukaryotic cells, but has also led directly to the cloning of related genes from

more complex eukaryotes.

II. Gene Transfer in Plants and Animals

Although the cells of complex eukaryotes are not amenable to the simple genetic

manipulation possible in yeasts, gene function can still be assayed by the introduction of cloned DNA

into plant and animal cells. Such experiments, generally called gene transfer, have proven critical to

addressing a wide variety of questions, including studies of the mechanisms that regulate gene

expression and subsequent protein processing. In addition, gene transfer has enabled the identification


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and characterization of genes that control animal cell growth and differentiation, including a variety of

genes responsible for abnormal growth of human cancer cells.

The method for introducing DNA into animal cells was initially developed for infectious viral

DNA and is therefore called transfection (a portmanteau derived from the words transformation and

infection). DNA can be introduced into animal cells in culture by one of a number of methods,

including direct microinjection into the cell nucleus, coprecipitation of DNA with calcium phosphate

to form small particles that are taken up by the cells, incorporation of DNA into lipid vesicles called

liposomes that fuse with the plasma membrane, and exposure of cells to brief electrical pulses that

transiently open pores in the cellular plasma membrane in a process known as electroporation. The

DNA taken up by most cells is transported to the nucleus, where it can be transcribed for several days;

this is a phenomenon known as transient expression. In a smaller fraction of cells (usually less than

1%), the foreign DNA becomes stably integrated into the cell genome and is transferred to progeny

cells at cell division, just as with any other cellular gene. These stably transformed cells can be isolated

if the transfected DNA contains a selectable marker, such as resistance to a drug that inhibits the

growth of normal cells. Thus, any cloned gene can be introduced into mammalian cells by being

transferred along with a drug resistance marker that can be used to isolate stable transformants. The

effects of such cloned genes on cell behavior – for example, cell growth and differentiation – can then

be analyzed.

Animal viruses can also be used as vectors for more efficient introduction of cloned DNA into

cells. Retroviruses are particularly useful in this respect, since their life cycles involve the stable

integration of DNA into the genome of infected cells. Consequently, retroviral vectors can be used to

efficiently introduce cloned genes into a wide variety of cell types, making them an important vehicle

for a broad array of applications.

Cloned genes can also be introduced into the germ line of multicellular organisms, allowing

them to be studied in the context of the intact animal rather than in cultured cells. This is shown in

Figure 37.14. One method used to produce mice that carry such transgenic, or foreign, genes is the

direct microinjection of cloned DNA into the pronucleus of a fertilized egg. The injected eggs are then

transferred to foster mothers and allowed to develop to term. In a fraction of the progeny (usually less

than 10%), the foreign DNA will have integrated into the genome of the fertilized egg and is therefore

present in all cells of the animal. Since the foreign DNA exists in both the germ and somatic cells of

the animal, it is transferred by breeding to new progeny.

III. Embryonic Stem Cells

Embryonic stem (ES) cells provide an alternative means of introducing cloned genes into

mice. ES cells can be established in culture from early mouse embryos. They can also be reintroduced


825

into early embryos, where they participate normally in development and can give rise to cells in all

tissues of the mouse, including germ cells. It is thus possible to introduce cloned DNA into ES cells in

culture, select stably transformed cells, and then introduce those cells back into mouse embryos. Such

embryos give rise to chimeric offspring in which some cells are derived from the normal embryo cells

and some from the transfected ES cells. In some such mice, the transfected ES calls are incorporated

into the germ line. Breeding these mice therefore leads to the direct inheritance of the transfected gene

by their progeny. This is shown, along with the pronucleus method of producing transgenic animals,

in Figure 37.14.

Figure 37.14: Introduction of genes to produce transgenic mice. Microinjection method

(bottom): DNA is microinjected into one of the two pronuclei of a fertilized mouse egg

(fertilized eggs contain two pronuclei, one from the egg and one from the sperm). The

microinjected eggs are then transferred to foster mothers and allowed to develop. Some

of the offspring are transgenic, meaning that they have incorporated the DNA into their

genome. Embryonic stem cell method (top): ES cells are cultured cells derived from early

mouse embryos (blastocysts). DNA can be introduced into these cells in culture, and

stably transformed ES cells can be isolated. These transformed ES cells can then be

injected into a recipient blastocyst, where they are able to participate in normal

development of the embryo. Some of the progeny mice that develop after transfer of


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injected embryos to foster mothers therefore contain cells derived from transformed ES

cells, as well as from the normal cells of the blastocyst. Since these mice are a mixture of

two different cell types, they are referred to as chimeric. Offspring carrying the

transfected gene can be produced by the breeding of chimeric mice in which descendants

of the transformed ES cells have been incorporated into the germ line.

III. Gene Transfer in Plants

Cloned DNA can also be introduced into plant cells. One approach is to bombard plant cells

with DNA-coated microprojectiles, such as small particles of tungsten. The DNA-coated particles are

projected directly into the plant cells; some of the cells are killed by the effects of the impact, but others

survive and become stably transformed. In addition, the tumor inducing (Ti) plasmid, taken from the

bacterium Agrobacterium tumefaciens, provides a novel vehicle for the introduction of cloned DNA into

many species of plants. In nature, Agrobacterium attaches to the leaves of plants, and the Ti plasmid is

transferred into plant cells where it becomes incorporated into sensitive cells of the host. Since many

plants can be regenerated from single cultured cells, transgenic plants can be established directly from

the cells into which recombinant DNA has been introduced in culture. This procedure is much

simpler than the production of transgenic animals! Indeed, many economically important types of

plants, including tomatoes, soybeans, corn, and potatoes are transgenic varieties.

IV. Mutagenesis of Cloned DNAs

In classical genetic studies, as with those conducted using bacteria or yeasts, mutants are the

key to identifying genes and understanding their function. In such studies, mutant genes are detected

because they result in observable phenotypic changes – for example, temperature-sensitive growth or a

specific nutritional requirement. The isolation of genes by recombinant DNA, however, has opened a

different approach to mutagenesis. It is now possible to introduce any desired alteration into a cloned

gene and to determine the effect of the mutation on gene function. Such procedures have been called

reverse genetics, since a mutation is introduced into a gene first and its functional consequence is

determined later. Introducing mutations into cloned DNA is called in vitro mutagenesis.

Cloned genes can be altered by many in vitro mutagenesis procedures, which can lead

to the introduction of deletions, insertions or single nucleotide alterations. The most common method

of mutagenesis is the use of synthetic oligonucleotides to generate nucleotide changes in a DNA

sequence. In this procedure, a synthetic nucleotide bearing the desired mutation is used as a primer for

DNA synthesis. Newly synthesized DNA molecules containing the mutation can then be isolated and


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characterized. For example, specific amino acids of a protein can be altered in order to characterize

their role in protein function.

Variations of this approach, combined with the versatility of other methods for

manipulating recombinant DNA molecules, can be used to introduce virtually any desired alteration

in a cloned gene. The effects of such mutations on gene expression and function can then be

determined by introduction of the gene into an appropriate cell type. In vitro mutagenesis has thus

allowed detailed characterization of the functional roles of both the regulatory and protein-coding

sequences of cloned genes.

V. Introducing Mutations into Cellular Genes

Although the transfer of cloned genes into cells, particularly in combination with in vitro

mutagenesis, provides a powerful approach to studying gene structure and function, such experiments

fall short of defining the role of an unknown gene in a cell or intact organism. The cells used as

recipients in the transfer of cloned genes usually have normal copies of the gene in their chromosomal

DNAs already; after transfer, these normal copies continue to perform their roles in the cell.

Determining the biological role of a cloned gene therefore requires that the activity of the normal

cellular gene copies be eliminated. Several approaches can be used to either inactivate the

chromosomal copies of a cloned gene or inhibit normal gene function, both in cultured cells and in

transgenic mice.

Mutating chromosomal genes is based on the ability of a cloned gene introduced into

a cell to undergo homologous recombination with its chromosomal copy. In homologous

recombination, the cloned gene replaces the normal allele, so mutations introduced into the cloned

gene in vitro become incorporated into the chromosomal copy of the gene. In the simplest case,

mutations that inactivate the clone gene can be introduced in place of the normal gene copy in order

to determine that gene’s role in cellular processes.

Recombination between transferred DNA and the homologous chromosomal gene

occurs frequently in yeast, but is a rare event in mammalian cells. Thus, inactivation of mammalian

cells by this approach is technically difficult. Possibly because the genomes of mammalian cells are so

much larger than those of yeasts, most transfected DNA that integrates into the recipient cell genome

does so at random sites by recombination with unrelated sequences. However, various procedures

have been developed to both increase the frequency of homologous recombination and to select and

isolate the transformed cells in which homologous recombination has occurred. It is feasible to

inactivate any desired gene in mammalian cells by this approach. Importantly, genes can be readily

inactivated in mouse embryonic stem cells, which can then be used to generate transgenic mice. These

mice can be bred to yield progeny containing mutated copies of the targeted gene on both homologous


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chromosomes, so the effects of inactivation of a gene can be investigated in the context of the intact

animal. In addition, cells can be cultured from mouse embryos containing the mutated gene copies, so

the functions of target genes can also be studied in cell culture. The biological activities of thousands of

mouse genes have been investigated in this way, and such studies have been critically important in

revealing the roles of many genes in murine development.

Homologous recombination has been used to systematically inactivate, or knock out,

every gene in yeast. This resulted in a collection of genome-wide yeast mutants that is available for

scientists to use to study the function of any desired gene. Methods also now exist to conditionally

knock out genes in specific mouse tissues, allowing the function of a gene to be studied in a defined cell

type (for example, in a nerve or liver cell) rather than in all cells of the organism.

VI. Interfering with Cellular Gene Expression

As an alternative to gene inactivation by homologous recombination, a variety of approaches

can be used to specifically interfere with gene expression or function. One method that has been used

to inhibit expression of a desired target gene is the introduction of antisense nucleic acids into cultured

cells. RNA or ssDNA complementary to the mRNA of the gene of interest (referred to as the antisense

gene) hybridizes with mRNA and blocks its translation into protein. Moreover, the RNA-DNA

hybrids resulting from the introduction of antisense DNA molecules are usually degraded within the

cell. Antisense RNAs can be introduced directly into cells, or cells can be transfected with vectors that

have been engineered to express antisense RNA. Antisense DNA is usually in the form of short

oligonucleotides, which can either be transfected into cells, or in many cases, taken up by cells directly

from the culture medium.

Recently, RNA interference (RNAi) has emerged as an extremely effective and widely used

method for interfering with gene expression at the level of mRNA. As discussed in a previous chapter,

RNAi is a major regulatory mechanism used by cells to control expression at both the transcriptional

and translation level. When double-stranded RNAs are introduced into cells, they are cleaved into

short double-stranded molecules by the enzyme Dicer. These short double-stranded molecules, called

short interfering RNAs (siRNAs), then associate with a complex of proteins known as the RNA-

induced silencing complex (RISC). Within this complex, the two strands of siRNA separate and the

strand complementary to the mRNA (the antisense strand) guides the complex to the target mRNA by

complementary base pairing. The mRNA is then cleaved by one of the RISC proteins. The RISC-

siRNA complex is released following degradation of the mRNA and can continue to participate in

multiple rounds of mRNA cleavage, leading to effective destruction of the targeted mRNA.

RNAi has been established as a potent method for interfering with gene expression in C.

elegans, Drosophila, Arabidopsis, and mammalian cells, and provides a relatively straightforward approach


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to investigating the function of any gene whose sequence has previously been established. In addition,

libraries of double-stranded RNAs or siRNAs that cover a large fraction of genes in the genome are

used to screen C. elegans, Drosophila, and human cells to identify novel genes involved in specific

biological functions, such as cell growth or survival.

In addition to inactivating a gene or inducing degradation of an mRNA, it is sometimes

possible to interfere with the function of proteins within cells. One approach is to microinject

antibodies that block the activity of the protein against which they are directed. Alternatively, some

mutant proteins interfere with the function of their normal counterparts when they are expressed

within the same cell. (For example, they may compete with the normal protein for binding to its target

molecule.) Cloned DNAs encoding such mutant proteins (called dominant inhibitory mutants) can be

introduced into cells by gene transfer and used to study the effects of blocking normal gene function.


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Chapter 37 Problems


Human embryonic stem cells (hES) have the capacity for pluripotency. As cells differentiate

from the embryonic stem cell state, this capacity is lost. Two experiments were conducted in order to

investigate whether a differentiated adult nucleus can de-differentiate, giving rise to a fully

reprogrammed nucleus.

Experiment 1

Three new populations of cells were created from a sheep embryo, a sheep fetus and the

mammary gland of an adult female sheep, or ewe. Nuclear contents from each of the three donor cell

populations were transferred into enucleated unfertilized eggs from a ewe in a process referred to as

somatic cell nuclear transfer (SCNT), and then implanted in recipient ewes. Prior to SCNT, donor

cells were induced to exit the growth phase and enter quiescence. The nuclear content donors and

recipients were different breeds. Eight ewes gave birth to live lambs; each lamb shared the

morphological characteristics of the breed from which the donors cells were derived, not that of the

recipient ewe. The scientists’ findings are summarized in Table 1.

Experiment 2

Specific diploid hES cells transfected to express genes for green fluorescent protein (HUES6-

GFP cells) were mixed with diploid human BJ fibroblast (BJ) cells containing a drug-resistance marker

for puromycin. The cells were cultured and colonies bearing one of two distinct morphologies

appeared. Each colony was screened for successfully fused hybrid (HUES6-GFP/BJ) cells. Once

selected, HUES6-GFP/BJ cells, which displayed an appearance consistent with that of (hES) cells,

were isolated and their DNA content analyzed.

Injection of HUES6-GFP/BJ cells into mice led to the formation of embryoid bodies (EBs)

and teratomas. Immunostaining revealed that both contained neuroectoderm-derived βIII-tubulin,

mesoderm-derived myosin, and endoderm-derived alpha-fetoprotein proteins. Introduction of a

transgenic reporter gene revealed that embryonic genes were reactivated in the HUES6-GFP/BJ cells.

Genome-wide transcriptional profiling demonstrated that expression of genes active exclusively in BJ

cells were suppressed in HUES6-GFP/BJ cells, despite the fact that DNA content analysis indicated

that no genetic information was lost in the fusion process.


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Table 1: Development of sheep embryos reconstructed from one of three different donor

cell types.

No. of fused

couplets (%)

No. recovered

from oviduct (%)

No. of morula/

blastocyst (%)

No. of

pregnancies/no.

of recipients (%)

No. of

live

lambs

Mammary

epithelium

277 (63.8) 247 (89.2) 29 (11.7) 1/13 (7.7) 1

Fetal

fibroblast

172 (84.7) 124 (86.7) 34 (27.4)/13 (54.2) 4/10 (40) 3

Embryo-

derived

385 (92.9) 231 (85.3) 90 (39)/36 (39) 14/27 (51.8) 4

1. Based on the results of the experiments described in the passage, which of the following statements

is most justifiable?

A. Differentiated cells in all stages of the cell cycle can be reprogrammed to an embryonic-like state

by transfer of nuclear contents into oocytes or by fusion with embryonic stem cells.

B. The introduction of transcription factors found exclusively in embryonic stem cells can induce

somatic cells to revert to an embryonic-like state.

C. Viable adult organisms capable of reproduction can be produced from the transfer of nuclear

contents from embryonic cells into oocytes.

D. Differentiation does not involve irreversible modification of cells that prevents their

reprogramming to the embryonic state.

2. In Experiment 1, in what stage of the cell cycle were the mammary epithelial cells immediately

prior to their introduction into the enucleated cell?

A. G0

B. G2

C. S

D. M


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3. Genes associated with puromycin resistance were introduced into the fibroblasts used in

Experiment 1 via transduction. Which of the following methods of gene transfer is another example of

transduction?

A. A harmless strain of the bacteria Streptococcus pneumoniae made virulent after exposure to a heat-

killed virulent strain

B. Transfer of genes for tetracycline resistance between Shigella and E. coli bacteria following direct

contact between them

C. The integration of a plasmid conferring kanamycin resistance by the bacteria Pseudomonas via a

DNA pump

D. A new genotype arising in recombinant Salmonella typhimurium strains due to the action of a

bacteriophage

4. Mesoderm-derived myosin was found during a histological examination of the teratoma sample

from Experiment 2. Which of the following structures in humans contains tissues also of mesodermal

origin?

A. retina

B. spinal cord

C. lung bronchi

D. ventricles of the heart

5. DNA content analysis was performed on successfully fused hybrid HUES6-GFP/BJ cells. How

many chromosomes, in terms of the human haploid number n, did the cells contain?

A. n

B. 2n

C. 4n

D. both 2n and 4n cells were observed


6. In electrophoresis, nucleic acids are separated on the basis of differences in their:

A. positive or negative charge only.

B. size only.


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C. ability to hybridize to the stationary phase.

D. positive or negative charge and size.

7. The size of the human genome is approximately 3 x 106 kb. If the size of an insert for a BAC

vector is between 120-300 kb, then what is the maximum number of BAC clones required to produce

a genomic library of human DNA?

A. 1,000

B. 10,000

C. 25,000

D. 40,000

8. If E. coli is grown for several generations in media containing 15N, before the cells are transferred to

a media containing only 14N and grown for two additional generations, what will be the ratio of the

number of bacterial cells containing only 14N to cells containing both 14N and 15N?

A. 1:4

B. 1:2

C. 1:1

D. 2:1

9. How would you expect dactinomycin, an inhibitor of DNA-directed RNA synthesis, to affect

replication of the non-retroviral RNA virus, influenza?

A. Transcription of viral RNA would cease.

B. Translation of virally-encoded proteins would be affected.

C. The rate of viral entry into cells would increase.

D. There would be no change in viral replication.

10. What feature of a cloning vector would allow for the isolation of stably transfected mammalian

cells?

A. an endonuclease recognition sequence

B. a stable origin of replication

C. an antibody probe binding site

D. a selectable marker


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1. D.

Experiment 1 demonstrates that adult mammary epithelium cells, under the circumstances in which

they were used, may be returned to an embryonic-like state of pluripotency; only a pluripotent cell,

when introduced into an oocyte, would be capable of giving rise to all of the tissues present in the lamb

born from nuclear transfer between adult mammary epithelial cells and recipient oocytes. Experiment

2 demonstrates a similar finding. According to passage information, HUES6-GFP/BJ cells are

consistent in appearance and behavior with hES cells and do not carry out the transcriptional program

found in un-hybridized BJ fibroblast cells. This suggests that differentiated adult cells do not undergo a

process of irreversible modification. If they are able to give rise to all of the cell types in an organism,

they must reversibly return to a de-differentiated, embryonic-like state in order to do so. All of this is

consistent with choice D, the correct answer. According to passage information, cells from which

nuclear contents were donated in Experiment 1 were induced to exit the growth phase and enter one

of quiescence prior to the nuclear transfer. It cannot be definitively concluded whether cells at any

phase of the cell cycle will behave as these quiescent cells did. Choice A is false and an incorrect

answer. Neither experiment introduced transcription factors, making choice B incorrect as well. In

Experiment 1, viable lambs were born, and while those lambs may well survive into adulthood and be

capable of reproducing, neither experiment provides evidence directly supporting such a finding. This

makes choice C false and an incorrect answer.

2. A.

According to passage information, the cells from which nuclear contents were donated in Experiment

1 were induced to exit the growth phase and enter one of quiescence prior to the nuclear transfer. This

implies that the cells were in G0, a period entered into by certain cells from the G1 phase, in which

cells are neither dividing nor preparing to divide. Many cells, including those in nervous and cardiac

muscle tissue, will remain in a quiescent state permanently after terminally differentiating. Other

mature cell types, including renal and hepatic parenchymal cells, may be induced to re-enter the cell

cycle. Choice A is true and is the correct answer.

3. D.

Transduction is the process by which genetic information is transferred between bacteria by a virus.

This is what is occurring in choice D, wherein a bacteriophage (a virus which injects its genome into

the cytoplasm of a bacterial host) transfers genetic information between Salmonella bacteria. Choice D

is therefore our correct answer. Transfection refers to the introduction of nucleic acids into cells by


835

non-viral means. Both choices A and C are examples of transfection, which is sometimes also referred

to as transformation (particularly when the genetic information is introduced into non-animal

eukaryotic cells). Cells can be transfected by physical or chemical treatment or via a non-bacterial

biological carrier particle. Choice B is an example of bacterial conjugation – the transfer of genetic

material, usually in the form of a plasmid, between bacterial cells via indirect cell-to-cell contact or via

a pilus or pilus-like bridging structure.

4. D.

The mesoderm, ectoderm and endoderm are the three primary germ layers of the developing human

embryo. The mesoderm gives rise to smooth, cardiac and skeletal muscles, connective tissue and bone,

the endothelium of blood vessels, red and white blood cells, the kidneys and the adrenal cortex.

Therefore choice D is the correct answer. Tissue of the central and peripheral nervous system,

epidermis and hair, and the sensory structures of the eye arise from the ectoderm, making choice A

and B incorrect. The endoderm gives rise to nearly all of the GI tract and the cells lining the glandular

organs which are associated with it (including the pancreas and liver), respiratory tract as well as

portions of the bladder, thymus, thyroid and urethra. As a result, choice C is incorrect.

5. C.

Both the HUES6-GFP cells and BJ fibroblast cells that contributed to the fusion product were somatic

cells that should have contained the human diploid number of 46 (2n) chromosomes. The passage

indicates that no genetic information was lost in Experiment 2 during the formation of the HUES6-

GFP/BJ cells, meaning that 92 (4n) chromosomes should be detected in the analysis performed.

Choice C is the correct answer, while choices A, B and D are false.

6. B.

Due to their phosphate backbone, nucleic acids are negatively charged. During electrophoresis, species

are separated on the basis of their respective sizes, as reflected by their respective electrophoretic

mobilities. This makes choice B correct. When attempting DNA electrophoresis, the stationary phase

commonly used is agarose gel. When agarose cools, it solidifies as the individual molecules interact.

These interactions form a three-dimensional mesh structure with small pores that DNA can fit

through. The DNA has to find its way through the agarose. Smaller particles migrate faster than

larger ones and reach the positive anode first.


836

7. C.

Dividing the size of the human genome, by the smallest possible insert size indicates that at most (3 x

106)/(1.2 x 102) = 2.5 x 104 or 25,000 BAC clones would be needed to cover the entire genome.

8. C.

If E. coli is grown for several generations in media containing 15N, then when transferred to media

containing only 14N, all cells will contain only 15N. Following the semi-conservative method of DNA

replication, after one generation of growth, all cells will contain both 14N and 15N; after two

generations, half of the cells will contain only 14N and the other half will contain both 14N and 15N.

The ratio of the number of cells containing only 14N to cells containing both 14N and 15N is therefore

1:1.

9. D.

The question states that the influenza virus is non-retroviral, implying the virus replicates via RNA-

directed RNA synthesis. Therefore, addition of an inhibitor of DNA-directed RNA synthesis such as

dactinomycin would not impact viral replication.

10. D.

When developing a stable transfection, researchers use selectable markers to distinguish transient from

stable transfections. Co-expressing the marker with the gene of interest enables researchers to identify

and select for cells that have the new gene integrated into their genome while also selecting against the

transiently transfected cells. For example, a common selection method is to co-transfect the new gene

with another gene for antibiotic resistance and then treat the transfected cells with a specific antibiotic

for selection. Only stably transfected cells with resistance to the antibiotic will survive in long-term

cultures, allowing for the selection and expansion of the desired cells. The hallmark of stably

transfected cells is that the foreign gene becomes part of the genome and is therefore replicated.

Descendants of these transfected cells, therefore, will also express the new gene, resulting in a stably

transfected cell line. As a result, only choice D is correct.

control of gene expression in...

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