Chapter 2. Basic Principles of Molecular Biology and Genomics

 

Chapter 2. Basic Principles of Molecular Biology and Genomics

BASIC PRINCIPLES OF MOLECULAR BIOLOGY AND GENOMICS: INTRODUCTION

In June 2000, it was announced that both a corporate effort and a government consortium had succeeded in

sequencing all of the human genome. This was followed by the publication of that sequence in February 2001

(Lander et al. 2001; Venter et al. 2001). For anyone involved in biology or medicine, these events represented a

revolution in the technical and conceptual approach to both research and therapy.

It seems that humans are far less complex than most scientists had previously thought. Rather than having

100,000–150,000 genes, as was once the belief, humans may have only about 30,000 genes. At this point, not all of

those genes have an identified function, and it is becoming clear that many gene products have more than one

function. Perhaps more importantly, genes that have been identified in a single cell type may have an entirely

different function in other cell types. Some other genes may enjoy a brief and transient expression during the

process of embryonic development only to play an entirely different role in the adult. Truly understanding those

genes and gene products will revolutionize all of science, and this may be especially true for psychiatry.

Consider that prior to identification of the genome, psychiatric genomics has been limited to studies of

chromosomal linkage wherein a putative gene for a disorder could be roughly localized to a given region of a

chromosome. The burgeoning understanding of the human genome now occurring has led to a rudimentary

understanding of genetic variation among humans. In many humans, a single base or single nucleotide is modified,

and it is a combination of knowing the entire genetic code and determining aberrations in individuals with disease

that will allow the pinpointing of specific genes associated with psychiatric diseases.

New advances and technology are also furthering our understanding of the genome. Microarrays, which permit one

to put several genes on a chip, show the ability of a given cell or tissue to activate given genes. Sometimes, during

a disease process, inappropriate genes are activated or inactivated. Identification of these genes also helps to shed

light on the disease process and on possible therapy. At the same time that rapid advances are being made in

understanding the genome, rapid advances in molecular biology are allowing the manipulation of genes and

proteins in individual nerve cells. The development of molecular and cellular models for neuropsychiatric disease

has also permitted tremendous advancement in our understanding of both biochemical defects and possible new

approaches toward ameliorating those defects.

In this chapter, we present information about genetics, genomics, and the genome and explain modern molecular

biology and the investigative methods used in that field. We also discuss pathophysiology, as related to

neuropsychiatry and molecular strategy, and introduce findings from studies on the cell biology of the neuron that

help us to understand both psychopharmacology and the biology of the brain and mind.

CELL BIOLOGY OF THE NEURON

To appreciate the molecular biology presented in this chapter, it is necessary to describe the components of the

neuron that process signals that directly or indirectly modify the aspects of the genome described below.

Neurons—specialized cells that function to transmit signals to other neurons, muscles, and secretory cells—contain

four basic domains (Figure 2–1), and these domains serve to receive signals, process and integrate signals, conduct

impulses, and release transmitter.

FIGURE 2–1. Diagram of a typical neuron.Print: Chapter 2. Basic Principles of Molecular Biology and Genomics http://www.psychiatryonline.com/popup.aspx?aID=416283&print=yes…

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As described in the text, this neuron is divided into zones for the reception of signals (input = dendrites), integration of signals

(regulation = nucleus and soma), conduction of signals (axon), and transmission of signals (axon terminal).

The nucleus resides in the cell body (the signal processing domain) and contains the DNA that codes for the genes

expressed by neurons. Activation of a given gene results in the generation of a messenger RNA (mRNA), which is

then translated into a protein (see below). Although such events are common to all cells, neural cells are unique in

some aspects of molecular signaling. Of importance, the variety of gene expression is far greater in the brain than

in any other organ or tissue. Some estimates are that in aggregate, the brain expresses as much as 10 times the

number of genes expressed in any other tissue. This does not mean that individual neurons undergo a much greater

gene expression. Rather, it suggests an extraordinary heterogeneity among neurons and glia, which allows for a

rich regulation when those neurons and glia assemble into the elaborate network of the human brain.

mRNA molecules exported from the nucleus are translated into proteins by ribosomes in the endoplasmic reticulum.

Note that most of the protein production occurs in the cell body, although there is also some mRNA in the dendrites

(Steward and Wallace 1995). This means that newly made proteins must be transported from that cell body to the

axon terminal, a distance as great as 1 m. These proteins are often packaged in vesicles, and specialized “motor”

molecules transport packaged proteins down microtubule “tracks” at the cost of adenosine triphosphate (ATP)

hydrolysis (Setou et al. 2000).

ESSENTIAL PRINCIPLES OF GENE EXPRESSION

Genes and DNA

The DNA double helix transmits genetic information from generation to generation and is the repository of

information required to guide an organism’s development and interaction with the environment. The role of DNA in

storing and transferring hereditary information depends on the innate properties of its four constituent bases.

There are two purine bases, adenine (A) and guanine (G), and two pyrimidine bases, cytosine (C) and thymine (T).

Within the DNA double helix, A is complementary to T, and G is complementary to C. Each block of DNA that codes

for a single RNA or protein is called a gene, and the entire set of genes in a cell, organelle, or virus forms its

genome. Cells and organelles may contain more than one copy of their genome. There are 46 chromosomes in a

typical human cell; when “unraveled,” the total DNA of a single cell is approximately 1 m in length. The 46 human

chromosomes consist of 22 pairs of autosomes and 2 sex chromosomes, either XX for females or XY for males. Such

a large amount of genetic material is effectively packaged into a cell nucleus, which is also the site of DNA

replication and transcription. Only a small percentage of chromosomal DNA in the human genome is responsible for

encoding the genes that act as a template for RNA strands; there are approximately 20,000–25,000 genes total, of

which about 10,000–15,000 genes are expressed in any individual cell.

Among RNA strands, only ribosomal RNA (rRNA), transfer RNA (tRNA), and small nuclear RNA (snRNA) have

independent cellular functions. Most cellular RNA, mRNA, serves as a template for protein synthesis. RNA, like DNA,

is also composed of four nucleotide building blocks. However, in RNA, the nucleotide uracil (U) takes the place of

thymine (T), and RNA is a flexible single strand that is free to fold into a variety of conformations. Thus, the

functional versatility of RNA greatly exceeds that of DNA.

Chromosomal DNA contains both genes and more extensive intergenic regions. Some regions of DNA in genes act

as the template for RNA, but some regions are responsible for regulatory functions. The distribution of genes on

chromosomes is not uniform: some chromosomal regions, and indeed whole chromosomes, are richly endowed with

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conferred by the nucleotide sequence of a DNA molecule is referred to as cis-regulation, because the regulatory and

transcribed regions occur on the same DNA molecule. cis-Regulatory elements that determine the transcription

start site of a gene are called basal (or core) promoters; other cis-regulatory elements are responsible for tethering

different activators and repressor proteins to DNA. There are specific regions of DNA that bind to regulatory

proteins. These regulatory proteins may be encoded at any regions in the genome, and because they are not coded

by the stretch of DNA to which they bind, they are sometimes called trans-acting factors. Trans-acting factors that

regulate the transcription of DNA are also called transcription factors.

DNA Replication

Chromosomal DNA must be replicated to coordinate with cell division. Replication begins at a sequence called the

origin of replication. It involves the separation of the double helix DNA strands over a short length and the binding

of enzymes, including DNA and RNA polymerases. During DNA replication, each existing strand of DNA serves as a

template for the synthesis of a new double helix that contains one old strand and one strand that is newly

synthesized but complementary. This process is known as semiconservative replication. In the process of cell

division, each of the 46 double helices is replicated and folded into chromosomes.

Transcription

Only a fraction of all the genes in a genome are expressed in a given cell or at a given time. These genes undergo

the process of transcription, in which an RNA molecule complementary to one of the gene’s DNA strands is

synthesized in a 5′ to 3′ direction, using nucleotide triphosphates. Transcription can be classified into three discrete

steps: initiation, mRNA chain elongation, and chain termination. Transcriptional regulation may occur at any step in

the process; however, initiation appears to be the primary control point because, in a sense, it is the rate-limiting

step. Localization of the transcription start site and regulation of the rate of transcription are essential to initiation.

The cis- and trans-acting factors described above all regulate the initiation of transcription.

Translation

Each mRNA in a cell can code for the primary amino acid sequence of a protein, using a triplet of nucleotides

(codon) to represent each of the amino acids. Some amino acids are represented by more than one codon, because

there are more triplet codons than there are amino acids. The codons in mRNA do not interact directly with the

amino acids they specify. The translation of the individual codons of mRNA into protein depends on the presence of

another RNA molecule, tRNA, which has a cloverleaf structure. On the top leaf of the tRNA structure, three

nucleotides form a complementary codon (an anticodon) to each mRNA nucleotide triplet. Thus, each mRNA

nucleotide triplet can code for a specific amino acid. Each tRNA carries an amino acid corresponding to its

anticodon, and when thus “charged,” the complex is termed aminoacyl-tRNA. Anticodons of aminoacyl-tRNA bind

with mRNA codons in ribosomes. Ribosomes, a complex of rRNA and enzymes needed for translation, provide the

structure on which tRNA can bind with the codons of mRNA in sequential order.

Initiation of protein synthesis involves the assembly of the components of the translation system. These

components include the two ribosomal subunits, the mRNA to be translated, the aminoacyl-tRNA specified by the

first codon in the message, guanosine triphosphate (GTP), and initiation factors that facilitate the assembly of this

initiation complex. In eukaryotes, there are at least 12 distinct translation initiation factors (Roll-Mecak et al.

2000). After the ribosome recognizes the specific start site on the mRNA sequence, which is always the codon AUG

coding for methionine, it slides along the mRNA molecule strand and translates the nucleotide sequence one codon

at a time, adding amino acids to the growing end of the polypeptide chain (the elongation process). During

elongation, the ribosome moves from the 5′-end to the 3′-end of the mRNA that is being translated. The binding of

GTP to the elongation factor tu (EFtu) promotes the binding of aminoacyl-tRNA to the ribosome (Wieden et al.

2002). When the ribosome finds a stop codon (UAA, UGA, or UAG) in the message RNA, the mRNA, the tRNA, and

the newly synthesized protein are released from the ribosomes, with the help of release factors that also bind GTP.

The translation process is stopped, and a nascent protein exists.

It is noteworthy that initiation, elongation, and release factors undergo a conformational change upon the binding

of GTP. In this regard, they are similar to the G proteins (both heterotrimeric G proteins and small “ras-like” G

proteins) involved in cellular signaling (Halliday et al. 1984; Kaziro et al. 1991).

REGULATION OF GENE EXPRESSION

Chromatin and DNA Methylation

Biophysics and molecular biology have revealed that chromatin consists of a repetitive nucleoprotein complex, the

nucleosome. This particle consists of a histone octamer, with two copies of each of the histones (H2A, H2B, H3, and

H4), wrapped by 147 base pairs of DNA. In the octamer, histones H3 and H4 are assembled in a tetramer, which is

flanked by two H2A–H2B dimers. A variable length of DNA completes the second turn around the histone octamer

and interacts with a fifth histone, H1. H2A, H2B, H3, and H4 are variously modified at their amino- and

carboxyl-terminal tails to influence the dynamics of chromatin structure and function (Ballestar and Esteller 2002;Print: Chapter 2. Basic Principles of Molecular Biology and Genomics http://www.psychiatryonline.com/popup.aspx?aID=416283&print=yes…

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Keshet et al. 1986; Kornberg and Lorch 1999; Strahl and Allis 2000). Although chromatin provides structure to

chromosomes, it also plays a critical role in transcriptional regulation in eukaryotes because it can repress gene

expression by inhibiting the ability of transcription factors to access DNA. In fact, chromatin ensures that genes are

inactive until their expression is commanded. In the activation process, cells must attenuate nucleosome-mediated

repression of an appropriate subset of genes by means of activator proteins that modify chromatin structure. An

activator protein displaces nucleosomes, which permits a complex of proteins (general transcription factors) to

bind DNA at a promoter and to recruit RNA polymerase.

Cytosine methylation at CpG dinucleotides is the most common modification of the eukaryotic genome. In

vertebrates, methylation occurs globally throughout the genome, with the exception of CpG islands. These are

CG-rich regions of DNA that stretch for an average of ~1 kilobase (kb), coincident with the promoters of 60% of

human RNA polymerase II–transcribed genes. Methylation of cytosines at CpG represses transcription (Ballestar

and Esteller 2002). Genetic imprinting, a process by which particular paternal or maternal genes are inactivated

throughout a species, is at least partly controlled by DNA methylation. Specific proteins binding to methylated DNA

may establish a bridge between chromatin and DNA methylation. They recruit histone deacetylases (HDACs) to

activate a methylated promoter, which in turn deacetylates histones, leading to a repressed state (Ballestar and

Esteller 2002; Keshet et al. 1986).

RNA Polymerases

There are three distinct classes of RNA polymerase—RNA polymerase I, RNA polymerase II, and RNA polymerase

III—in the nucleus of eukaryotic cells, and they are designed to carry out transcription. RNA polymerase I

synthesizes large rRNA molecules. RNA polymerase II is mainly used to yield mRNA and, subsequently, proteins.

RNA polymerase III produces snRNA, small rRNA, and tRNA molecules. Each class of RNA polymerase recognizes

particular types of genes. However, RNA polymerases do not bind to DNA directly. Rather, they are recruited to DNA

by other proteins that bind to promoters (Figure 2–2).

FIGURE 2–2. Transcription factors and RNA polymerase II complex.

Typical transcription factors contain DNA-binding domains, protein dimerization domains, and transcription activation domains.

Some transcription factors (e.g., cAMP response element–binding protein [CREB]) may be modified by phosphorylation. The

transcription activation domain interacts with an RNA polymerase II (Pol II) complex to induce transcription. TATA binding

protein (TBP) binds to the TATA box element and associates with general transcription factors (TFII). This gene transcription

apparatus recruits Pol II to the appropriate gene.

mRNA is transcribed from DNA by RNA polymerase II with heterogeneous nuclear RNA (hnRNA), an intermediate

product. The core promoter recognized by RNA polymerase II is the TATA box (Hogness box), a sequence rich in

nucleotides A and T, which is usually located 25–30 bases upstream of the transcription start site. The TATA box

determines the start site of transcription and orients the basal transcription complex that binds to DNA and recruits

RNA polymerase II to the TATA box; thus, it establishes the 5′ to 3′ direction in which RNA polymerase II

synthesizes RNA. The formation of the basal transcription complex is promoted by a TATA binding protein (TBP)

that binds to a core promoter, together with multiple TBP-associated factors and other general transcription

factors. Enhancers are DNA sequences that increase the rate of initiation of transcription by RNA polymerase II

through its interaction with transcription factors, which can be located “upstream” or “downstream” of the

transcription start site. Enhancer elements are important to cell-specific and stimulus-dependent expression of

hnRNA. Some RNA polymerase II species, including those for many genes that are expressed in neurons, lack a

TATA box and possess instead an initiator, a poorly conserved genetic promoter element.

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Transcription factors act as the key regulators of gene expression. Sequence-specific transcription factors typically

contain physically distinct functional domains (see Figure 2–2). Numerous transcription factors have been found.

Some of them translocate to the nucleus to bind their cis-regulatory elements in response to activation reaction,

such as nuclear factor B (NF- B). However, some transcription factors are already bound to their cognate

cis-regulatory elements in the nucleus under basal conditions and are converted into transcriptional activators by

phosphorylation. cAMP (cyclic 3′-5′-adenosine monophosphate) response element–binding protein (CREB), for

example, is bound to regions of DNA, called cAMP response elements (CREs), before cell stimulation. CREB can

promote transcription when it is phosphorylated on a serine residue (ser133), because phosphorylated CREB can

interact with a coactivator, CREB-binding protein, which in turn contacts and activates the basal transcription

complex. Of interest, CREB-binding protein possesses intrinsic histone acetyltransferase (HAT) activity. The activity

of most transcription factors is regulated through second-messenger pathways. CREB can be activated via

phosphorylation at ser133 by second messengers, such as cAMP, Ca++, and growth factors (Figure 2–3).

FIGURE 2–3. Activation of cAMP response element–binding protein (CREB) via different signal transduction

pathways.

Signal cascades are activated by external stimuli, such as hormones or neurotransmitters and growth factors. Arrows indicate

the interaction between pathways. AC = adenylyl cyclase; C = catalytic subunits of PKA; Ca++ = calcium; CaMK IV =

calmodulin-dependent kinase IV; cAMP = cyclic 3′-5′-adenosine monophosphate; CBP = CREB-binding protein; Epac =

exchange protein activated by cAMP; ERK = extracellular-regulated kinase; G s = subunit of the stimulatory G protein; P =

phosphorylation; PKA = cAMP–dependent protein kinase; R = regulatory subunits of PKA; Rap and Ras = small GTPases (small

proteins that bind to guanosine triphosphate [GTP]); RSK2 = ribosomal S6 kinase 2.

CREB is a molecule that is widely implicated in learning and memory in many species. Mice expressing mutant CREB

isoforms show impaired memory, but this is dependent on the genetic background of the mice (Graves et al. 2002).

CREB-binding protein (CBP) is a transcriptional coactivator with CREB. A partial-knockout mouse model, in which

CBP activity is lost, exhibits learning deficiencies (Oike et al. 1999). Further, Rubinstein-Taybi syndrome (RTS) is an

autosomal-dominant dysmorphic syndrome that results in severe impairment of learning and memory. The RTS

gene has been mapped to chromosome 16 and identified as CBP. The loss of function of CBP is likely one important

contributing factor to the learning and memory defects seen in RTS (Murata et al. 2001; Oike et al. 1999; Petrij et

1. 1995).

Posttranscriptional Modification of RNA

The mRNA of prokaryotes can be used without any modification to direct protein synthesis, but posttranscription

processing of mRNA is needed in eukaryotes. The DNA sequences that code for mRNA (exons) are frequently

interrupted by intervening DNA sequences (introns). When a protein-coding gene is first transcribed, the hnRNA

contains both exons and introns. Before the transcript exits the nucleus, its introns are removed and its exons arePrint: Chapter 2. Basic Principles of Molecular Biology and Genomics http://www.psychiatryonline.com/popup.aspx?aID=416283&print=yes…

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spliced to form mature mRNA (Figure 2–4). The hnRNA that is synthesized by RNA polymerase has a

7-methyl-guanosine “cap” added at the 5′ end. The cap appears to facilitate the initiation of translation and to help

stabilize the mRNA. In addition, most eukaryotic mRNA has a chain of 40–200 adenine nucleotides attached to the

3′ end of the mRNA. The poly (A) tail is not transcribed from DNA; rather, it is added after transcription by the

nuclear enzyme poly (A) polymerase. The poly (A) tail may help stabilize the mRNA and facilitate mRNA exit from

the nucleus. After the mRNA enters the cytoplasm, the poly (A) tail is gradually shortened.

FIGURE 2–4. Transcription and RNA splicing.

The horizontal black line between exons indicates an intron. The region before the first exon is the 5′ regulatory region of the

gene, such as a TATA box. There also are cis-regulatory elements in introns and downstream of the last exon. The

heterogeneous nuclear RNA (hnRNA), containing both exons and introns, is spliced to form mRNA. mRNAs are then exported

from the nucleus to the cytoplasm, where they will direct the synthesis of distinct proteins.

RNA Editing

RNA editing has been detected in eukaryotes ranging from single-celled protozoa to mammals and plants and is

now recognized as a type of RNA process (posttranscriptional modification of RNA) that differs from the established

processes of RNA splicing, 5′ end formation, and 3′ endonucleolytic cleavage and polyadenylation (DeCerbo and

Carmichael 2005; Kable et al. 1997). The conversion of adenosine to inosine was observed first in yeast tRNA

(Grosjean et al. 1996) but has since been detected in viral RNA transcripts and mammalian cellular RNA (Bass

1997; Simpson and Emeson 1996). The inosine residues generated from adenosines can alter the coding

information of the transcripts, as inosine is synonymous for guanosine during transcript translation. For example,

upon A-to-I editing, the CAC codon for histamine is transformed to CIC, coding for arginine. RNA editing can have

dramatic consequences for the expression of genetic information, and in a number of cases it has been shown to

lead to the expression of proteins not only with altered amino acid sequences from those predicted from the DNA

sequence but also with altered biological functions (Bass 2002; Burns et al. 1997).

The enzymes for RNA editing are referred to as adenosine deaminases that act on RNA (ADARs). ADARs target RNA

that is double-stranded and convert adenosines to inosines by catalyzing a hydrolytic deamination at the adenine

base (Bass 2002). Mammals have several ADARs, of which two (ADAR1 andADAR2) are expressed in most tissues

of the body (Seeburg and Hartner 2003). RNA editing may also catalyze the conversion of one or a few adenosines

in a transcript to inosines (Maas et al. 1997; Stuart and Panigrahi 2002). On the other hand, RNA editing can

convert numerous adenosines to inosines in RNA. This type of editing is thought to be the result of aberrant

production of dsRNA (DeCerbo and Carmichael 2005) and has been suggested to lead to RNA degradation (Scadden

and Smith 2001), nuclear retention (Zhang and Carmichael 2001), or even gene silencing (Wang et al. 2005).

The 5-HT2C receptor is a G protein–coupled receptor that is well known to have variants generated by A-to-I editing

(Burns et al. 1997). 5-HT2C receptor transcripts can be edited at up to five sites, potentially generating 24 different

receptor versions, and hence a diverse receptor population. The RNA-edited 5-HT2C receptor affects ligand affinity

and the efficacy of G protein coupling (Berg et al. 2001; Wang et al. 2000; Yang et al. 2004). The unedited form of

the 5-HT2C receptor has the highest affinity to serotonin and exhibits constitutive activity independent of serotonin

levels. When RNA is edited, the basal activity of the 5-HT2C receptor is suppressed, and agonist potency and efficacy

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Modification of the Nascent Polypeptide Chain

The posttranslational modifications described above occur after translation is initiated. They may include removal

of part of the translated sequence or the covalent addition of one or more chemical groups that are required for

protein activity. Some of these modifications, such as glycosylation or prenylation, represent an obligatory step in

the synthesis of the “finished” protein product. In addition, many proteins may be activated or inactivated by the

covalent attachment of a variety of chemical groups. Phosphorylation, glycosylation, hydroxylation, and prenylation

are common types of covalent alterations in posttranslational modifications. A number of different enzymes

coordinate these processes, and they represent a major portion of the events of cellular signaling.

APPROACHES TO DETERMINING AND MANIPULATING GENE EXPRESSION

Changes in gene expression within the central nervous system (CNS) have profound effects on all other aspects of

the organism. Changes in gene expression are causally associated not only with the development of the CNS but

also with the complex phenomena of brain function, such as memory formation, learning, cognition, and affective

state. Changes in gene expression likely underlie the pathogenesis of many sporadic or inherited CNS-related

disorders, such as Alzheimer’s disease, Huntington’s disease, depression, and schizophrenia. Thus, insight into and

characterization of gene expression profiles are necessary steps for understanding how the brain functions at the

molecular level and how malfunction will result in disease. Molecular biological and genomic technologies such as

gene mapping and cloning, DNA libraries, gene transfection and expression, and gene knockout and gene targeting

have provided numerous benefits to neuropsychopharmacology. Genomic methods applied to pedigree and

population samples of patients with psychiatric disorders may soon make it possible to identify genes contributing

to the etiology and pathogenesis of these diseases and to provide a potential basis for new therapies.

Cloning of DNA

The cloning of DNA confers the ability to replicate and amplify individual pieces of genes. Cloning can be performed

with genomic DNA or complementary DNA (cDNA). cDNA is synthesized artificially from mRNA in vitro with the aid

of reverse transcriptase. Cloned genomic DNA may contain any stretch of DNA, either introns or exons, whereas

cloned cDNA consists only of exons. For cloning (see Figure 2–5 for outline of the process), the desired pieces of

DNA (often called “inserts”) are connected with the DNA of genetically engineered vectors or plasmids, and the

vectors are introduced into hosts, such as bacteria or mammalian cells. A DNA library is a collection of cloned

restriction fragments of the DNA of an organism that consists of random pieces of genomic DNA (i.e., genomic

library) or cDNA (i.e., cDNA library). Complete cDNA libraries contain all of the mRNA molecules expressed in a

certain tissue. Sometimes cDNA libraries can be made from a specific tissue in a distinct circumstance. For example,

a cDNA library could be made from cerebral cortex in rats undergoing transient forebrain ischemia (Abe et al.

1993).

FIGURE 2–5. Outline of gene cloning.Print: Chapter 2. Basic Principles of Molecular Biology and Genomics http://www.psychiatryonline.com/popup.aspx?aID=416283&print=yes…

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See text for details.

The cloning of disease-related genes is an important step toward insight into the pathogenesis of diseases and

development of new drugs against these diseases. If a protein is suspected of involvement in pathogenesis, a

nucleic acid sequence can be deduced from the partial amino acid sequence. cDNA libraries are screened with the

partial sequence in order to fish out a complete clone of interest. The isolated cDNA molecules are used to infect

bacteria, which amplify the cloned cDNA. Bacteria expressing these cDNA molecules will often make the protein of

interest, which can be detected with antibody. A variation on the above technique uses antibodies against the

purified proteins to screen a DNA library transfected into bacteriophages. The bacteriophages containing the “right”

cDNA can be retrieved, and the sequence of cDNA can be analyzed.

Polymerase Chain Reaction

Polymerase chain reaction (PCR) is a rapid procedure for in vitro enzymatic amplification of specific segments of

DNA. Amplification of the genes of interest occurs by selecting and synthesizing “primers”—stretches of DNA that

span the region of interest to be “filled in”—and by heating DNA to make it single stranded, allowing polymerase

and primers to bind. The polymerase then reads the “blank” stretch of DNA between the primers and synthesizes

DNA corresponding to the region of interest. This is usually done in “thermal cyclers” that heat the DNA at regular

intervals and allow “cycles” of PCR to amplify the genes repetitively. To avoid continual addition of polymerase, the

DNA polymerase used is often from bacteria that inhabit hot springs or hot ocean vents. This polymerase is not

denatured by heating and can therefore support several cycles of PCR.

A variation on PCR is reverse transcriptase PCR (or RT-PCR), in which the RNA is the template. Reverse

transcriptase (the “RT” of RT-PCR) is employed to synthesize cDNA from the RNA. Enzymatic amplification of this

cDNA is then accomplished with PCR.

Real-time PCR is based on the method of RT-PCR, following the reverse transcription of RNA into cDNA. Real-time

PCR requires suitable detection chemistries to report the presence of PCR products, as well as an instrument to

monitor the amplification in real time through recording of the change in fluorescence (Wittwer et al. 1997).Print: Chapter 2. Basic Principles of Molecular Biology and Genomics http://www.psychiatryonline.com/popup.aspx?aID=416283&print=yes…

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Generally, chemistries in PCR consist of fluorescent probes. Several probes exist, including DNA-binding dyes like

EtBr or SYBR green I, hydrolysis probes (5′-nuclease probes), and hybridization probes (Valasek and Repa 2005).

Each type of probe has its own unique characteristics, but the strategy for each is to link a change in fluorescence

to amplification of DNA (Kubista et al. 2006; Lind et al. 2006). The instrumentation to detect the production of PCR

must be able to input energy for excitation of fluorescent chemistries at the desired wavelength and simultaneously

detect a particular emission wavelength. Many instrument platforms are available for real-time PCR. The major

differences among them are the excitation and emission wavelengths that are available, speed, and the number of

reactions that can be run in parallel (Kubista 2004).

Positional Cloning

Disease genes can sometimes be isolated with the aid of positional cloning, a process also known as reverse

genetics (Collins 1995). Positional cloning is the process used to identify a disease gene, based only on knowledge

of its chromosomal location, without any knowledge of the biological function of the gene. Positional cloning

requires a genetic map of unique DNA segments or genes (genetic markers), with known chromosomal location,

that exist in several alternative forms (alleles). These allelic variations (polymorphisms) allow comparisons of the

wild type as the “diseased” genotype.

Linkage analysis is a method of localizing one or more genes influencing a trait to specific chromosomal regions.

This is performed by examining the cosegregation of the phenotype of interest with genetic markers. Relatives who

are phenotypically alike will share common alleles at markers surrounding the genes influencing the phenotype,

whereas other relatives (i.e., those who are phenotypically dissimilar) will not carry these alleles. To carry out

linkage analyses, investigators need, minimally, a set of families in which phenotyped individuals have known

relationships to one another and the genotypes of these individuals, including one or more genetic markers.

Once the chromosomal location of the disease gene has been ascertained, the area of chromosomal DNA can be

cloned. Until recently, the process of positional cloning involved laborious efforts to build a physical map and to

sequence the region. (The sequencing of the human genome has obviated this step.) Physical maps can be

produced by isolating and linking together yeast and/or bacterial artificial chromosomes containing segments of

human DNA from the region. These fragments are then sequenced and ordered, and from these data, the genomic

DNA sequence for the region of the candidate gene region is determined.

Differential Display

The technique of differential display is designed to determine the complement of genes being expressed (mRNAs)

by a tissue or an organ at a given point in time. The establishment of differential display is dependent on the

random amplification and subsequent size separation of cDNA molecules (Liang et al. 1992). With RT-PCR

amplification with specific oligo-T primers (one- or two-base anchored oligo-dT primers, such as oligo-T-XC,

oligo-T-XG, oligo-T-XT, and oligo-T-XA; X = G/A/C), four separate cDNA synthesis reactions are performed. These

cDNA synthesis reactions form the four pools of cDNA for one original mRNA population. The resulting cDNA

molecules from each RT-PCR reaction are amplified, using the same primer of the reverse transcription step plus

randomly chosen primers. Because the randomly chosen primers will anneal at various locations upstream of the

oligo-T site, many individual cDNA fragments of different sizes are amplified in each PCR reaction. cDNA fragments

derived from different original mRNA populations are sized and then separated on parallel gels to analyze the

presence of unique bands. The differentially expressed cDNA fragments can be excised from gels, cloned, and

further characterized by a variety of technologies based on different purposes, such as in situ hybridization,

sequences, and Northern blot analysis.

Differential display is a useful tool in identifying region-specific mRNA transcripts in brain. The molecular markers

for these regions can be found by screening for gene expression in specific brain regions or nuclei (Mizushima et al.

2000; Tochitani et al. 2001). In addition, under different stimulation or behavioral conditions, the changes in gene

expression can be explored by differential display (Hong et al. 2002; Q. R. Liu et al. 2002; Mello et al. 1997; Tsai et

1. 2002). This technique has even been adapted to indicate the RNA expression profile of an individual neuron

(Eberwine et al. 1992). Many genes related to ischemia or Alzheimer’s disease in the CNS were also isolated by

means of differential display (Doyu et al. 2001; Imaizumi et al. 1997; Tanaka et al. 2002). Genes that are activated

in response to chronic drug treatment (e.g., opiates or antidepressants) can also be identified this way.

Furthermore, as described later in this chapter (see “Microarray Technology”), streamlined technologies (gene

arrays) are now available for this purpose.

Gene Delivery Into Mammalian Cells

The introduction of recombinant DNA (including desired genes) into cells is becoming an increasingly important

strategy for understanding certain gene functions in neurons or glia. These advances in gene function studies have

benefited the diagnosis of, and therapy for, a variety of disorders that affect the CNS. The delivery of desired genes

into cells may also provide the technological base for insight into molecular mechanisms of brain function and,

ultimately, for gene therapy in the CNS.Print: Chapter 2. Basic Principles of Molecular Biology and Genomics http://www.psychiatryonline.com/popup.aspx?aID=416283&print=yes…

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Vectors and Delivery

For gene transfer, it is necessary to incorporate the desired cDNA into various vectors, such as appropriate

plasmids or replication-deficient viruses. Vectors used for transfection generally possess two essentially

independent functions: 1) carrying genes to the target cell and 2) expressing the genes properly in the target cell.

There are currently many commercial vectors with different features from which to choose, and the choice is

dependent on the purpose of the experiments and on the characterizations of exogenous desired genes. In one

form of transfection, stable transfection, cells expressing the gene of interest can be actively selected by a marker

(e.g., neomycin resistance genes, Neo), and the cDNA or other type of foreign DNA is stably integrated into the DNA

of a chromosome. In transient transfection, however, cells express the gene of interest for a few days. Cells for

hosting the foreign DNA can be either established cell lines or primary cultured neuronal cells. The desired cDNA

delivered into cells may be native genes, fragments of the genes, mutant genes, or chimeric genes.

The main barrier to the delivery of DNA into cells is getting the foreign DNA through the cellular membrane. Over

the past decades, various methods have been developed to convey the foreign DNA molecules into mammalian

cells. These include chemical or physical techniques, such as calcium phosphate coprecipitation of DNA, liposome

fusion, electroporation, microinjection, ballistic injection, and viral infection. At present, methods dependent on

incorporation of DNA into cationic liposomes (e.g., lipofectin) are used most widely. These methods of transfection

are accessible to both cell lines and cultured primary neuronal cells. The ratio of DNA to liposomal suspension, cell

density, and time duration of exposure to the DNA–liposomal complex must be optimized for each cell type in

culture.

Viral Vectors

Over the past few years, several viral vectors with low toxicity, high infection rate, and persistent expression have

extended the methodology of delivery of genes to mammalian cells. These viruses include DNA viruses, such as

adenoviruses and adeno-associated viruses, herpes simplex viruses, and RNA retroviruses. Recently, as a result of

advances in genetic manipulation, adenoviruses and adeno-associated viruses have been more widely applied to

gene transfer. The advantages of adenoviruses are 1) the ability to carry large sequences of foreign DNA, 2) the

ability to infect a broad range of cell types, and 3) an almost 100% expression of the foreign gene in cells.

Human adenovirus is a large DNA virus (36 kb of DNA) composed of early genes (from E1 to E4) and later genes (L1

to L5). Wild-type adenovirus cannot be applied to gene transfer, because it causes a lytic infection. Thus,

recombinant adenoviruses with defects of some essential viral genes are used for gene delivery. These recently

developed adenoviral expression systems are safe; such systems have the capacity for large DNA inserts and allow

for relatively simple adenoviral production (Harding et al. 1997; He et al. 1998).

The process of gene transfer into cells (cell lines and primary cells) via recombinant adenoviruses is simple, but the

optimal viral titer, the time of exposure to virus, and the multiplicity of infection should be optimized for each cell

type. Cell lines and a variety of primary neuronal cells have been infected by adenoviruses (Barkats et al. 1996;

Chen and Lambert 2000; Hughes et al. 2002; Koshimizu et al. 2002; Slack et al. 1996). In addition, recombinant

adenoviruses containing the desired genes can be delivered to neurons in vivo via intracerebral injection into

particular brain areas (Bemelmans et al. 1999; Benraiss et al. 2001; Berry et al. 2001; Neve 1993).

Identification of Ectopic Gene Expression

After delivery of the desired genes into cells, identification of their proper expression is necessary. The general

strategies for identification (Chalfie et al. 1994; Kohara et al. 2001; Yu and Rasenick 2002; Yu et al. 2002b) include

The measurement of some functional changes elicited by gene expression in targeted cells. If an enzyme is expressed,

this would involve measuring the activity of that enzyme.

1.  

The detection of the proteins coded by the desired genes by techniques relying on antibodies, such as western blot,

immunoprecipitation, and immunocytochemistry. Expression of the desired genes tagged with some epitopes, such as HA,

GST, and His-tag, can also be determined with antibodies directed against these epitopes.

2.  

The use of green fluorescent protein (GFP) as an indicator. GFP, a protein from the jellyfish Aequorea victoria, can be

either cotransfected with the desired genes or fused with the desired genes before incorporation into cells. The

fluorescence from GFP is easy to detect using fluorescent microscopy, and this allows the monitoring of gene expression in

living cells. Furthermore, if GFP is fused with the gene of interest, its fluorescence provides a useful tool for studying the

function and cellular localization of proteins coded by the genes of interest.

3.  

At this point, several different “colors” have been developed through mutation of the initial GFP gene (Shaner et al.

2005). Depending on the wavelengths of excitation, they can be used to localize multiple protein species, or

fluorescence resonance energy transfer (FRET) can be used to demonstrate that two target proteins are in close

(<10 nm) proximity. FRET uses two fluors with little spectral overlap and depends on the emission of one of the

fluors exciting the other.

INHIBITION OF CELLULAR GENE EXPRESSIONPrint: Chapter 2. Basic Principles of Molecular Biology and Genomics http://www.psychiatryonline.com/popup.aspx?aID=416283&print=yes…

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RNA Interference

The previous edition of this textbook (2004) mentioned RNA interference as a new technique with great potential

for simple manipulation of gene expression. In September 2006, Andrew Z. Fire and Craig C. Mello were awarded a

Nobel Prize for this versatile discovery. RNA interference (RNAi) is believed to be a biologically conserved function

in a wide range of eukaryotic species. It may play a role in protection against double-stranded RNA viruses (Sijen et

1. 2001) and genome-invading transposable elements (Provost et al. 2002; Volpe et al. 2002). Triggered by dsRNA,

RNAi identifies and destroys the mRNA that shares homology with the dsRNA. Thus, the expression of a particular

gene can be suppressed by introducing dsRNA whose antisense strand sequence matches the mRNA sequence. Fire

et al. (1998) first described RNAi in the nematode Caenorhabditis elegans as sequence-specific gene silencing in

response to double-stranded RNA. The mechanism of RNAi is partly understood, and key proteins involved in the

pathway have been identified. In brief, the basic process of RNAi involves the following steps. In a first initiation

step, Dicer, an enzyme of the RNase III family, initiates ATP-dependent fragmentation of long dsRNA into 21- to

25-nucleotide double-stranded fragments, termed small interfering RNAs (siRNAs). These siRNAs are specifically

characterized by overhanging 3′ ends of two nucleotides and phosphorylated 5′ ends. The siRNA duplexes bind with

Dicer, which facilitates the formation of an siRNA/multiprotein complex called RISC loading complex (RLC). The

siRNA duplex in RLC then unwinds (which requires the protein Ago2) to form an active RNA-induced silencing

complex (RISC) that contains a single-stranded RNA (called the guide strand). The RISC recognizes the target RNA

through Watson–Crick base pairing with the guide strand and cleaves the target RNA. Finally, the RISC releases its

cleaved product and goes on to catalyze a new cycle of target recognition and cleavage (Figure 2–6) (Tomari and

Zamore 2005; Xia et al. 2005).

FIGURE 2–6. Schematic of the mechanism of RNA interference (RNAi) posttranscriptional knockdown of a gene

product.

The procedure starts with introduction (transfection, electroporation, or injection) of double-stranded RNA (dsRNA) or small

interfering RNA (siRNA) into cells, or expression of small hairpin RNA (shRNA) in cells with vectors encoding shRNAs. The

cellular ribonuclease (RNase) Dicer recognizes the long dsRNA molecules and shRNA. Subsequently the dsRNA is cleaved,

resulting in 21-nt RNA duplexes, the siRNAs. These siRNA molecules are then incorporated into the RNA-induced silencing

complex (RISC) multiprotein complex, where they are unwound by an adenosine triphosphate (ATP)–dependent process,

transforming the complex into an active state. Activated RISC uses one strand of the RNA as a bait to bind homologous RNA

molecules. The target RNA is cleaved and degraded, resulting in gene silencing.

RNAi Knockdown of Gene Expression

Knockdown of gene expression is accomplished by designing siRNA sequences that target the coding and noncoding

regions of the candidate mRNAs with perfect complementarity. In mammalian cells, administration of siRNAs is

effective in short-term investigations. Introduction of siRNA into cells is accomplished by transfection or

electroporation of either the specific siRNA itself or small hairpin RNA (shRNA) in which the hair loop is rapidly

cleaved to produce siRNA. Most of the proposed applications of RNAi incorporate 21-nt siRNA duplexes with 2-nt 3’Print: Chapter 2. Basic Principles of Molecular Biology and Genomics http://www.psychiatryonline.com/popup.aspx?aID=416283&print=yes…

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overhangs, which after chemical synthesis allow large-scale and uniform production of siRNA molecules that are

also amenable to stabilizing chemical modifications. Several commercial entities involved in the manufacturing of

siRNAs provide effective design algorithms online, which are based on a combination of mRNA target sequence and

secondary structures. In some siRNA–target combinations, the use of longer dsRNAs can increase the potency of

RNAi (Kim et al. 2005; Siolas et al. 2005).

For stable longer-term suppression, a gene construct coding for the shRNA with a Pol III or a Pol II promoter can

be applied (Brummelkamp et al. 2002; Y. Shi 2003; Yu et al. 2002a). RNA Pol II and III promoters are used to drive

expression of shRNA constructs (Amarzguioui et al. 2005), depending on the type of expression required. Pol III

promoters drive high levels of constitutive shRNA expression, and their transcription initiation points and

termination signals are well defined. Pol II promoter–driven shRNAs can be expressed tissue-specifically (Zeng et

1. 2002). Expressed shRNAs are efficiently incorporated into RISC, leading to a more potent inhibition of target

gene expression. Generally, viral vectors such as adenovirus, lentivirus, or adeno-associated virus can carry the

inhibitory construct into cells to achieve adequate expression for knockdown (Kim and Rossi 2007).

RNAi Applications

RNAi is a straightforward method for inducing sequence-specific silencing of one or more genes of interest with the

introduction of siRNAs. It has been a powerful tool to investigate gene function. Studies using RNAi have been

performed in neuronal cells to examine the functional roles of individual genes in developing growth cones and

neurite outgrowth (Eriksson et al. 2007; Hengst et al. 2006; J. Liu et al. 2007; Schmitz et al. 2007; Yanaka et al.

2007), ion channels (Geng et al. 2004; Lauver et al. 2006; Tahiliani et al. 2007), apoptosis (Yano et al. 2007; Zhang

et al. 2007), and a variety of signaling pathways (Meuer et al. 2007; Sanada and Tsai 2005; G. X. Shi et al. 2006;

Yamada et al. 2005). Furthermore, RNAi is now being used for the knockdown of gene expression in animals. It

appears to apply in virtually all mammalian species, as exemplified by its capability for silencing genes in mice, rats,

and goats (Peng et al. 2006; Zhou et al. 2007). Inducible RNAi based on the Cre-loxP system has also been

developed in transgenic mice, enabling the investigator to control gene silencing spatially and temporally (Chang et

1. 2004; Coumoul et al. 2005; Xia et al. 2006). Thus, it is possible to rapidly establish human genetic diseases as a

whole-animal model without the difficulty and expense of embryonic stem cell and gene targeting.

RNAi can knock down specific genes, making it evident that RNAi may be used to disrupt the expression of

disease-associated genes for therapeutic purposes. Recent findings have highlighted the possibility of RNAi as a

potential therapeutic approach to many human diseases, although a number of challenges need to be addressed,

such as the safety concerns and effective delivery of RNAi therapy. RNAi-based therapies for age-related macular

degeneration (AMD) and respiratory syncytial virus (RSV) have already reached clinical trials. Moreover, therapies

based on RNAi are also in preclinical development for other viral diseases (Rossi 2006), neurodegenerative

disorders (Raoul et al. 2006; Xia et al. 2005), and cancers (Pai et al. 2006).

RNAi has become a frequently used tool in a wide range of biomedical research. It has provided a convenient

method to study gene function via application of RNAi to cell lines, cultures, and embryonic stem cells. It is

anticipated that new uses for this tool will continue to proliferate.

Antisense Oligonucleotides

Antisense oligonucleotides (AOs) are short sequences of nucleotide with about 20 bases that are complementary to

some regions of the specific mRNA sequence of interest. When cells take up the short DNA molecules, synthesis of

the protein from the specific mRNA is inhibited. The mechanism by which AO blocks the specific protein production

remains unclear. Interfering with the selected mRNA stability or its translation may be one potential mechanism

(Phillips et al. 1996). Because of specificity, effectiveness, and reversibility, cell-based antisense experiments have

proven useful in studying gene and protein function.

PROTEOMIC TECHNIQUES FOR ALTERING CELLULAR FUNCTION

Not all manipulations of gene products are made at the gene level. It is possible to modify cell function by

interfering directly with gene products (i.e., the proteomic strategy). One mechanism for such manipulation

involves adding peptides that correspond to the active regions of proteins. These peptides bind to the molecular

targets of those proteins within the cell and block the downstream effects. Sometimes these peptides can mimic

the effect of the larger peptide. For example, a peptide corresponding to the carboxyl-terminal region of a G protein

(which interacts with G protein–coupled receptors) was used to block the interaction between receptors and G

proteins while “mimicking” the G protein to shift the receptor into a “high affinity” agonist-binding state (Rasenick

et al. 1994). Although this strategy usually involves adding peptides to cells made permeable with a detergent or

with electric current, it is also possible to incorporate DNA plasmids encoding peptides into cells (Gilchrist et al.

1999).

Another proteomic strategy for manipulating cellular processes involves expression of “dominant-negative”

proteins that are generated in sufficient amount to block the activity of the native protein in the cell. Usually this

strategy involves the construction of a mutant protein that is similar to the protein of interest but deficient at somePrint: Chapter 2. Basic Principles of Molecular Biology and Genomics http://www.psychiatryonline.com/popup.aspx?aID=416283&print=yes…

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active site. This inactive mutant protein, expressed in considerable excess over the native protein, competes with

the native protein for target sites and inhibits its activity (Osawa and Johnson 1991).

Proteomics and Beyond

Shotgun proteomics takes its name from shotgun DNA sequencing, in which long DNA sequences are disassembled

into shorter, easily readable components and reassembled. Shotgun proteomics works in much the same way as

the proteome of a cell is digested, subjected to analysis on mass spectrometry, and “reassembled” into proteins

identified through a sophisticated bioinformatic analysis of the protein “fingerprint.” At first glance, such an

approach might seem problematic, as a tripeptide would have the same mass regardless of the order of the amino

acids; however, ionization of the peptide reveals the order of the amino acids. It is the computational algorithms

that allow this identification. A recent review explains this in language that any biologist (or psychiatrist) can

understand (Marcotte 2007).

Analysis of the proteome has the potential to yield much more information than simply the identity of the proteins

being expressed in a given cell at a given time. Drugs that mimic, antagonize, or alter metabolism of

neurotransmitters and neuromodulators comprise the vast majority of the current psychiatric armamentarium.

These drugs, often used chronically, have been shown to have long-term effects, altering second-messenger

systems. These second messengers (e.g., cAMP, inositol-1,4,5-triphosphate [IP3], cyclic guanosine monophosphate

[cGMP]) activate or inhibit enzymes that modify proteins covalently. Phosphorylation, prenylation, and

glycosylation are examples of these modifications, all of which can be determined in mass spectroscopic proteomic

analysis.

In addition to—and, in part, due to—the modifications discussed above, molecules involved in neurotransmitter

response and responsiveness move among cellular compartments in response to neurotransmitters,

neuromodulators, and drugs that affect these. A simple example might be the agonist-induced depalmitoylation of

the subunit of the stimulatory G protein (Gs ) that results in its translocation from the plasma membrane

(Wedegaertner and Bourne 1994; Yu and Rasenick 2002). We have demonstrated that this translocation is due to

collection of G proteins in lipid rafts structures that either amplify or attenuate signaling (Allen et al. 2005, 2007). A

catalog of these protein shifts during disease or response to drug can be easily assembled.

As noted above, G protein–coupled receptors comprise targets for a major component of drugs used in psychiatry.

Curiously, many of these drugs were thought to have a single site of action until a screening/informatics approach

was applied to them. Receptoromics allows screening of compounds for their agonist or antagonist effects on

receptors measured through batteries of cells selectively expressing a single species of G protein–coupled receptor

(or other receptor or transporter). Results from these studies have been both informative and surprising (Strachan

et al. 2006). For example, atypical antipsychotics have been selected for their antagonist properties at the 5-HT2A

receptor. Some of these drugs, such as olanzapine, have been associated with excessive weight gain, and this

appears to be due to interaction with H1 receptors—not related to the therapeutic profile of the drug. Other

effective antipsychotic drugs may owe their efficacy to actions at a panel of receptors that, when combined,

contribute to a therapeutic whole. The ability to screen libraries of compounds against a large number of receptor

targets makes design and identification of new drugs an exciting possibility.

TRANSGENIC AND GENE-TARGETING TECHNIQUES

Over the past two decades, progress in the development of molecular genetic methods has enabled the

manipulation of genes in intact organisms, such as mice. The technologies have provided a powerful and useful tool

that allows the study of gene function and promotes understanding of the molecular mechanisms of disorders of

the brain and mind. The mouse genome is by far the most accessible mammalian genome for manipulation. Many

successful procedures for introducing new genes, expressing elevated levels of genes, and eliminating or altering

the function of identified target genes have been reported. Many mouse models produced by manipulating genes

may be used in a variety of fields relevant to neuroscience. It is noteworthy, however, that behavioral studies in

mice lag far behind those in rats. For this reason, genetically induced behavioral alterations in mice must be

interpreted with caution (Lucki et al. 2001).

Generally, transgenic mice are those expressing exogenous DNA because of the insertion of a gene into the mouse

genome. Usually that gene is randomly located within the mouse genome, often as several copies. The use of

transgenic mice has represented a major strategy for the investigation of genetic questions since the feasibility and

reproducibility of stably introducing DNA by microinjection into individual male mouse embryos were established

(Markert 1983). In DNA constructs used for the generation of transgenic mice, the gene of interest is located 3′ to

promoter sequences to produce a desired distribution of gene expression. The selection of the promoter is the most

important consideration in generating transgenic mice. Some promoters, such as platelet-derived growth factor

(PDGF), thy1 (a cell surface glycosylphosphatidylinositol-linked glycoprotein), prin (PrP), neuron-specific enolase

(NSE), and glial fibrillary acidic protein (GFAP), have demonstrated the ability to direct high-level expression of

exogenous genes in brain and/or in the neurons of mice (Hsiao et al. 1996; Kan et al. 2004; Nolte et al. 2001;

Sturchler-Pierrat et al. 1997). This level of expression can be modified by incorporation of a “tet-on” or “tet-off”Print: Chapter 2. Basic Principles of Molecular Biology and Genomics http://www.psychiatryonline.com/popup.aspx?aID=416283&print=yes…

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vector into the desired inserted gene. Depending on the nature of the switch (on or off), the mouse will express the

gene of interest when ingesting (or taken off) doxycycline.

“Gene targeting” refers to the homologous recombination that occurs between a specifically designed targeting

construct and the chromosomal target of interest, in which recombination at the target locus leads to replacement

of the native target sequence with construct sequence. The method enables the precise introduction of a mutant

into one of many murine genes and has proven invaluable in examining the roles of gene functions in complex

biological processes. Most of the target constructs are used to disrupt a target and to eliminate gene function

(conventional “knockout”). Generally, a gene-targeting construct that contains positive–negative selection markers

is prepared such that the target gene is interrupted by the gene for neomycin resistance, which also serves as a

positive selection marker, and a thymidine kinase (TK; Gusella et al. 1983) gene is adjacent to either one or both

ends of the homologous genomic sequence for negative selection. The positive–negative deletion scheme is

employed to enrich for homologous recombination.

The targeting construct is often introduced into mouse embryonic stem (ES) cells by electroporation. Cells that fail

to integrate the targeting construct into the genome are killed by application of neomycin in the culture medium

(positive selection). The majority of the remaining cells, in which the entire construct (including the TK gene)

inserts randomly, will die as a result of the incorporation of ganciclovir or fialuridine (inactive thymidine analogs),

which block DNA synthesis. Homologous recombinant clones that do not contain the TK gene are used to prepare

chimeric mice. Cells from these clones are microinjected into the fluid-filled cavity of 3.5-day-old embryos at the

blastocyst stage. The injected embryos are then surgically transferred into the uterus of pseudopregnant females.

These animals will give birth to chimeric mice. Breeding can be used to generate mice that are heterozygous and

homozygous for the mutation. Homozygous mutants may express the gene of interest in any cell of the body

(Figure 2–7).

FIGURE 2–7. Conventional gene disruption (“knockout”).

(A) Producing chimeric mice. First, a mutant allele is produced by replacing the coding exons of the desired gene with a

neomycin (neo) cassette and transferring it into embryonic stem (ES) cells. Second, genetically altered ES cells are

reintroduced into a developing blastocyst, where they contribute to the developing embryo. (B) Breeding chimeric mice.

When the germ cells of the resulting chimeric mouse (chimera) are ES cell–derived (germ-line mutation), the heterozygotesPrint: Chapter 2. Basic Principles of Molecular Biology and Genomics http://www.psychiatryonline.com/popup.aspx?aID=416283&print=yes…

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(+/–) can be produced by breeding. One-half of the offspring will be heterozygous. The heterozygous animals may be bred to

produce homozygous mice (–/–). TK = thymidine kinase; WT = wild type.

Use of Mutant Mice in Studies of Brain Disease

Transgenic mice produced by this method are generally gain-of-function mutants, because the transgene is

designed either to express a novel gene product or to disrupt a normal gene product by expressing a

“dominant-negative” alternative. It is also possible to put a DNA fragment in an opposite direction and hence to

produce transgenic mice expressing antisense RNA, which will reduce gene function (Jouvenceau et al. 1999;

Katsuki et al. 1988). Expression of mutant amyloid precursor protein (APP) or presenilin 1 (PS1) in mice has

generated many animal models that may be related to Alzheimer’s disease. These transgenic mice have facilitated

studies of the pathogenesis, molecular mechanisms, and behavioral abnormalities of Alzheimer’s disease

(Bornemann and Staufenbiel 2000; Janus et al. 2001). Nevertheless, because the transgene integrates into the

mouse genome randomly and often exists as several copies, the interpretation of studies with transgenic mice is

difficult.

Experiments with knockout mice have provided novel insights into the functional roles of neuronal genes and, in

some cases, animal models relevant to brain disorders. The targeted mutants in a gene of interest, however, can

sometime lead to embryonic lethality in mice, thus obscuring the particular role of that gene in a target tissue or in

the adult. Furthermore, in some instances, genes related to the gene that was eliminated undergo increased

expression in the knockout mice. In this case, the related gene compensates for the gene of interest and yields a

phenotype that resembles the “normal” animal. This has been seen for the knockouts of the axonal

microtubule-associated protein tau (Takei et al. 2000). Here, we take Huntington’s disease to exemplify the use of

transgenic mice in the study of neurological and psychiatric disease.

Huntington’s disease (HD) is a genetic neurological disorder that is inherited in an autosomal dominant manner.

The gradual atrophy of the striatum is its pathological hallmark. It has a prevalence of 3–10 affected subjects per

100,000 individuals in Western Europe and North America (Gil and Rego 2008). The first symptoms generally

appear in middle age, and the disease is progressive and invariably fatal 15–20 years after its onset (Ho et al.

2001). HD is caused by an expansion of cytosine-adenine-guanine (CAG) repeats in exon 1 of the HD gene. The HD

gene is located on the short arm of chromosome 4 (4p63) and encodes the protein huntingtin, composed of more

than 3,100 amino acids with a polyglutamine tail, which is widely expressed throughout the body in both neuronal

and nonneuronal cells.

The function of huntingtin has been revealed with different research approaches, especially those with transgenic

mice. Engineered knockout mice that disrupt exon 4 (Duyao et al. 1995), exon 5 (Nasir et al. 1995), or the promoter

(Zeitlin et al. 1995) of mouse gene homology, hdh, showed embryonic lethality. A subsequent study using mutant

human huntingtin to compensate for the absence of endogenous huntingtin rescued the embryonic lethality of mice

homozygous for a targeted disruption of the hdh gene (Leavitt et al. 2001). These suggest its essential role for

normal embryonic development. Moreover, condition knockout mice indicated that huntingtin is also required

throughout life, because adult mice are sterile, develop a progressive motor phenotype, and with short life span

after inactivating hdh gene during adulthood (Dragatsis et al. 2000). Furthermore, overexpression of wild-type

huntingtin, bearing 12 glutamine residues, in transgenic mice brought significant protection against apoptosis

triggered by NMDA (Leavitt et al. 2006), suggesting that huntingtin may play a role in cellular apoptosis.

As mentioned above, HD is a neurodegenerative disorder caused by uninterrupted CGA trinucleotide repeats that

located near the 5′-end in exon 1. Consequently, mutant huntingtin bears a string of consecutive glutamine

residues in the NH2-terminal, 17 amino acids downstream of the initiator (Gil and Rego 2008). The length of

glutamine residues in NH2-terminal of mutant huntingtin is the primary and predominant determinant for severity

of HD. To elucidate the mechanism of neurodegeneration in HD, multiple mouse models of HD have been

established. These models vary in terms of the site of transgene incorporation, promoter used, gene expression

levels, CAG repeat length, and background mouse strain used. These transgenic models have facilitated

investigations into potential pathogenic mechanisms of HD. At present, there are three categories of mouse model:

Mice expressing exon-1 fragments of human huntingtin gene (HD) containing polyglutamine mutations (R6/1, R6/2, and

R6/5). This transgenic mouse carries exon 1 of the HD gene with 115–155 CAG repeats. The transgene protein contains

the first 69 amino acids of huntingtin in addition to the number of residues encoded by the CAG repeats. Extensive

neuropathological analysis has been performed on the brains is R6/2 mice. The mice display subtle motor and learning

deficits at approximately 1 month and overt symptoms appear by 2 months, and they usually die at 3 or 4 months (Carter

et al. 1999; Lione et al. 1999; Murphy et al. 2000). It is worthy to note that the characteristic nuclear inclusions were first

detected with antibodies against the N-terminal portion of huntingtin in R6/2 mice. R6/2 mice show measurable deficits in

motor behavior that increase progressively until death. R6/2 mice are a model of HD to aim at studying the severity of

motor symptoms or the course of the disease. In R6/2 mice, many neurotransmitter receptors, such as NMDA, AMPA,

mGluR2, DA, and GABA, display abnormal response to their ligands (Ali and Levine 2006; Cepeda et al. 2004; Cha et al.

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1998; Dunah et al. 2002; Starling et al. 2005).

Mice expressing the full-length human HD gene. Yeast artificial chromosomes (YACs) were used to create a YAC

transgenic mouse model of HD that expresses the full-length human HD gene. The transgenic mice express human

transgenic huntingtin with 18, 46, 72, or 128 polyglutamine repeats. The YAC mice with 72 CGA repeats develop a

progressive motor phenotype, neuronal dysfunction, and selective striatal neurodegeneration similar to that seen in HD by

a 12-month timeline. YAC mice with 128 CAG repeats (YAC128 mice) exhibit initial hyperactivity, followed by the onset of

motor deficits and finally hypokinesis, which show phenotypic uniformity with low interanimal variability present. The

motor deficit in the YAC128 mice is highly correlated with striatal neuronal loss, providing a structural correlate for the

behavioral changes (Hodgson et al. 1999; Slow et al. 2003). These lines of transgenic mice may be extremely useful for

preclinical experimental therapeutics.

2.  

Knock-in HD transgenic mice. In knock-in mice, a mutated DNA sequence is exchanged for the endogenous sequence

without any other disruption of the gene. To establish the line of knock-in HD mice, CGA repeats in the murine hdh gene

were replaced with human mutant CAG repeats. These mice are characterized by a biphasic progression in behavioral

anomalies, and the nuclear inclusions appear late and are preceded by nuclear staining for huntingtin, followed by the

presence of microaggregates of the mutant protein in the nucleus and the neuropil (Menalled et al. 2002, 2003, 2005).

These knock-in mice are considered to be an ideal genetic model of HD to evaluate the effectiveness of new therapies and

to study the mechanisms involved in the neuropathology of HD.

3.  

The above examples illustrate both the potential and the current promise of manipulation of the mouse genome for

the study of human neuropsychiatric disease.

MICROARRAY TECHNOLOGY

Since the mid-1990s, a new and advanced microarray technology was developed that allows the study of gene

expression. This technology is proving to be a powerful research tool for gaining insight into the study of gene

expression and gene structure. The main large-scale application of the microarray is comparative expression

analysis. However, studies of DNA variation are also possible with microarray techniques, as are pharmacogenetics

applications.

For DNA microarrays, DNA sequences, DNA inserts of a library from PCR amplifications, cDNA clones, or synthetic

oligonucleotides can be immobilized on an impermeable rigid support (e.g., glass) in matrix spots. These

microarrays can be hybridized to labeled cDNA probes prepared from the mRNA extracted from the cell and tissue

of interest. The hybridization of the probe to each array component is measured to provide a quantitative measure

of the abundance of each array component in the probe. Currently, oligonucleotide-based DNA chips are generated

by the in situ synthesis of short (20- to 30-nucleotide) DNA fragments by either photolithography on a chip

(developed by Affymetrix, Santa Clara, CA) or ink-jet technology (developed by Rosetta Inpharmatics, Kirkland,

WA). The latter offers more speed in producing an array and increases the number of elements that can be arrayed

on a single chip. Hybridization to short oligonucleotides on a chip has a lower threshold of specificity than the older

hybridization techniques, but it is a more comprehensive and extremely rapid screening mechanism. The application

of DNA microarray technology has been used recently for characterizing gene expression profiles in human

diseases, such as multiple sclerosis (Whitney et al. 1999), cancers (Perou et al. 1999), and various

neurodegenerative diseases (Ginsberg et al. 2000).

In principle, any type of ligand-binding assay that relies on the product formation of an immobilized capture

molecule and a “binder” present in the surrounding solution can be miniaturized, parallelized, and performed in a

microarray format (Templin et al. 2002). Many microarray-based assays have emerged; these include studies of

DNA–protein interaction in a microarray format (Bulyk et al. 1999), enzyme–substrate arrays (Zhu et al. 2000), and

protein–protein binding assays (Zhu et al. 2001). Microarray-based technology is likely to accelerate basic research

in the area of molecular interactions and has the potential to change the diagnostic methods used for a variety of

human diseases.

CONCLUSION

Rapid advances in the identification of the human genome and in the methodology for genetic manipulation have

combined to open a window into the brain. We are accumulating knowledge of human gene mutations and their

connection to neurological and psychiatric diseases at a rapid pace. As genes are being identified, the proteins for

which they code are also becoming known. With this knowledge, the pathogenic mechanism of some diseases is

becoming apparent. Understanding these maladies at the molecular level is likely to lead to new methods of

diagnosis and novel approaches to therapy.

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