Rho-dependent termination and ATPases in transcript termination

Review

Rho-dependent termination and ATPases in transcript termination

John P. Richardson *

Department of Chemistry, Indiana University, 800 East Kirkwood Avenue, Bloomington, IN 47405, USA

Received 21 June 2002; accepted 21 June 2002

Abstract

Transcription factor Rho is a ring-shaped, homohexameric protein that causes transcript termination through actions on nascent RNAs that

are coupled to ATP hydrolysis. The Rho polypeptide has a distinct RNA-binding domain (RNA-BD) of known structure as well as an ATP-

binding domain (ATP-BD) for which a structure has been proposed based on homology modeling. A model is proposed in which Rho first

makes an interaction with a nascent RNA on a C-rich, primarily single-stranded rut region of the transcript as that region emerges from the

exit site of RNA polymerase. A subsequent step involves a temporary release of one subunit of the hexamer to allow the 3Vsegment of the

nascent transcript to enter the central channel of the Rho ring. Actions of the Rho structure in the channel on the 3Vsegment that are coupled

to ATP hydrolysis pull the RNA from its contacts with the template and RNA polymerase, thus causing termination of its synthesis.

D 2002 Elsevier Science B.V. All rights reserved.

Keywords: Rho factor; Transcript termination; Hexameric helicase; RNA-binding protein; NusG; Factor 2

1. Introduction

Rho factor is a protein used in bacteria for termination of

transcripts at certain sites. It serves an active role in

termination, employing the energy from ATP hydrolysis to

mediate the dissociation of a nascent transcript. In Escher-

ichia coli, Rho-dependent terminators are used at the ends of

genes and operons and at regulatory sites (attenuators)

preceding genes [1]. The RNA polymerase in bacteria can

also terminate transcription at other sites without the assis-

tance of Rho factor [1]. These intrinsic terminators are also

used at ends of genes and at attenuators. Both types of

terminators are used in E. coli and other bacteria for the

orderly expression of the genetic information.

RNA polymerases are well adapted to making complete

transcripts of genes. Once committed to the transcription of

a gene sequence, they form stable transcriptional elongation

complexes (TECs) with the DNA and nascent RNA. The

stability of the TEC is consistently high for most sequences,

but decreases greatly upon transcription of the DNA of an

intrinsic terminator [2–5]. A combination of features near

the 3V end of the nascent transcript and the interactions of

that part of the transcript with other components of the TEC

cause RNA polymerase to release its hold on the RNA, thus

allowing spontaneous release of the transcript. The sequen-

ces of an intrinsic terminator conform to a consensus, which

contains a 20 base pair stretch of DNA that encodes an RNA

that can form a stable stem-loop structure [6]. This sequence

is followed by seven to eight A residues in the template

strand. Release of the transcript occurs when the final seven

to eight U residues of the transcript are paired with the seven

to eight A residues of the template. A hybrid helix of rU

residues with dA residues is much less stable than hybrid

helices with other sequences [7]. This property, combined

with the formation of a stem-loop structure in the RNA in

the exit channel of RNA polymerase, is sufficient for the

release of the transcript to dissociate from the complex

before the next nucleotide is added.

Rho-dependent terminators are very different and are not

definable by a simple consensus sequence. Rho serves the

function of mediating release of the transcript at a DNA

sequence at which the transcription complex is too stable for

spontaneous release. Thus, a Rho-dependent terminator can

occur at sites where the sequence of an intrinsic terminator

would be incompatible with other functions of the sequence.

To mediate release, Rho binds to the RNA and uses

mechanical actions on the RNA that are coupled to ATP

hydrolysis to wrest the transcript from RNA polymerase and

the DNA template [8,9]. Rho is thus an ATP-driven machine.

The purpose of this review is to present the recent

advances in the structure and mechanism of Rho that have

0167-4781/02/$ - see front matter D 2002 Elsevier Science B.V. All rights reserved.

PII: S0167 -4781 (02 )00456 -6

* Tel.: +1-812-855-1520; fax: +1-812-855-8300.

E-mail address: [email protected] (J.P. Richardson).

www.bba-direct.com

Biochimica et Biophysica Acta 1577 (2002) 251–260

provided insights into how this machine terminates tran-

scription. Also, some aspects of the biology of the function

of Rho in bacteria will be considered, as will the properties

of some ATP hydrolysis-driven release factors for eukary-

otic RNA polymerases.

2. Structure

Rho functions as an assemblage of six identical, wedge-

shaped subunits arranged around a symmetry axis like the

segments of an orange [10]. The attempts that have been

made to get diffraction grade crystals of Rho protein have

not been successful. However, the morphology of the

protein has been determined by a three-dimensional recon-

struction from the electron micrographic images of nega-

tively stained protein adsorbed onto carbon grids [10]. The

images are primarily ring-shaped with six-fold symmetry.

Although some earlier studies suggested that Rho might

have D3 symmetry, these electron micrographic images

were more consistent with a six-fold or pseudo six-fold

rotational (C6) symmetry. The diameter of the rings at their

widest point was about 120 A. The height along the six-fold

axis was about 90 A.

In the EM analysis, most rings had six subunits but about

10% had only five subunits arranged in a gapped ring

structure. From sedimentation studies, it is known that Rho

readily dissociates into smaller assemblies at concentrations

below 1 AM [11,12], which is about the concentration in the

cell and just under the concentration used for the EM studies.

Thus, the gapped structure could be very relevant for Rho

function. In the current model for the mechanism of Rho

action, a segment of the RNA needs to be captured in the

center of a ring under conditions in which the protein would

not be able to thread onto a closed ring structure [13,14].

This could readily happen by capturing the RNA in the

center of the ring of a temporarily gapped structure.

A single subunit of E. coli Rho has 419 amino acid

residues [15]. An N-terminal segment consisting of the first

129 residues has been isolated and crystallized [16,17]. This

segment binds RNA by itself and is called the RNA-binding

domain (RNA-BD). The remaining part of Rho has a site for

binding ATP [18,19]. It has been termed the ATP binding

domain (ATP-BD), although it has not been isolated.

Molecular level structures of the free RNA-BD both in

crystals [17] and in solution [20] reveal that its first 50

residues are organized into an a-helical bundle while the

remaining part is a h-barrel having an OB fold. The OB fold

is a common structural motif in many proteins that bind

single-stranded polynucleotides. Other proteins with OB

folds are ribosomal protein S1, bacterial cold-shock proteins

and the single-stranded DNA-binding proteins. One of the

hallmarks of Rho function is a requirement for single-

stranded segments in the RNA. These single-stranded seg-

ments bind across the OB-fold segments of multiple sub-

units in the Rho hexamer.

Fig. 1. Views of two opposing subunits of the Rho hexamer. Two modeled subunits are arranged with rotational symmetry about the vertical axis with the plane

of the paper. The N-terminal 123 residues are represented by a black ribbon, while the C-terminal region (residues 126–419) is represented by a gray ribbon.

This model is based on the one described in Burgess and Richardson [23].

J.P. Richardson / Biochimica et Biophysica Acta 1577 (2002) 251–260252

To suggest a mechanism of Rho, it has been necessary to

use models for the structures of the ATP-BD and for the

organization of the protein into a hexamer. These models are

based on the known structure of the F1-ATPase, a protein

that is similar to Rho. When compared with all other known

protein sequences in the data base, the sequences that are

most closely similar to those of the Rho protein sequences

are those of the a and h subunits of the F1-ATPase [21].

These subunits are well conserved in a great variety of

organisms and are closely similar to each other. This

similarity is strongest for the ATP-BD of Rho and the

corresponding parts of those subunits. The identity in those

regions is about 21% and the similarity is about 43%. Rho

and the part of the F1-ATPase that contains the three a and

three h subunits are also morphologically very similar [10],

suggesting that the proteins have the same quaternary

organization. For that reason, the tertiary structures of the

a and h subunits can serve as models for the tertiary

structure of Rho, and the conglomerate of the three a and

three h subunits serve as a model for the quaternary

structure [22]. Molecular level models were made by

threading the aligned sequences of the ATP-BD of Rho

onto the coordinates of the corresponding parts of the a and

h subunits of bovine mitochondrial F1-ATPase followed by

an energy minimization to remove steric clashes [23]. The

known coordinates of Rho’s RNA-BD were then docked

onto the model for the six ATP-BDs, replacing the corre-

sponding parts of the a and h subunits at one pole of the

hexamer. The orientation for the RNA-BDs was suggested

from the three-dimensional reconstruction of the Rho struc-

ture from the EM images, to which the known structure of

the RNA-BD could be docked with a unique fit [10]. This

orientation has the clefts for binding RNA facing out toward

the periphery of the globular structure with the normals to

the face of the clefts at about a 45j angle to the six-fold axis.Fig. 1 is a diagram of the ribbon structures of two opposing

subunits.

3. Rho–RNA interactions

Rho factor is known to bind with relatively high affinity

to RNA [24,25] and to the transcription factors NusG

[26,27] and NusA [28]. The binding to RNA is essential

for Rho function and is responsible for the specificity of

Rho action. The interaction of Rho with RNA activates NTP

hydrolysis by Rho [29], a step that is essential for mediating

release of RNA transcripts from the transcription complex.

However, Rho can bind to RNA in the absence of an NTP

[24,25], and this interaction has been used to analyze the

specificity of the binding. Rho has a strong preference for

single-stranded RNA rich in C residues. Thus, it binds very

tightly to poly(C). Poly(C) becomes saturated with Rho at a

ratio of one Rho hexamer per 78 nucleotides or about 13

nucleotides per subunit [24,25]. When complexes of Rho

with poly(C) are treated with ribonuclease, an RNA segment

of about 78 residues is protected from digestion [24,30].

This result suggests that poly(C) is bound in a cleft that

extends continuously across the six subunits.

The interaction of RNA with individual subunits in the

Rho hexamer has been probed by measuring the binding of

oligo(C) molecules that are smaller than 13 nucleotides in

length. For rC10, Rho has three strong and three weak RNA-

binding sites that differ by about 10-fold in their affinities

for oligonucleotides [31,32]. This result indicates that not all

the subunits have identical conformations. The isolated

RNA-BD also binds oligo(C) [16]. However, it does so

with an affinity (the Kd for rC6 is about 2 AM) that is similar

to that for the weak binding sites in the Rho hexamer. Thus,

the organization of Rho into a hexameric structure may

entail a conformational change that puts half of the subunits

into a higher affinity form.

Both isolated RNA-BD and hexameric Rho bind to other

RNA oligonucleotides and RNA homopolymers but with

generally much weaker affinities. The affinity for the next

best RNA homo-oligonucleotide, oligo(U), is more than 10-

fold lower [32]. Both will also bind to single-stranded DNA

as well [16,24]. The isolated RNA-BD binds to oligo(dC)

with an affinity that is about five-fold lower than for the

corresponding oligo(rC) [33]. Thus, the site in the RNA-BD

has a strong preference for C residues and a slight prefer-

ence for RNA over DNA. A crystal structure of a complex

of rC9 with the isolated RNA-BD has revealed the reason

for the strong preference for C residues and for pyrimidines

in general [34]. In the complex, one cytosine base of one

residue makes three H-bond interactions with the side

chains of two key residues in the RNA-BD, two H-bonds

in a bi-dentate interaction with Arg-66 and the third with an

oxygen on the Asp-78 side chain. The base in the next

residue to the 5V side fits into a pocket that is only large

enough for a pyrimidine. Thus, at least two of the inter-

actions in one of the subunits are strongly cytosine or

pyrimidine specific. Evidence from NMR spectra of com-

plexes of oligonucleotides with the isolated RNA-BD indi-

cate that there is another C-specific interaction at residues

88 and 89, which extend from the short a-helix in the OB-

fold of the RNA-BD (T.K. Hitchins and G.S. Rule, personal

communication).

4. Attachment of Rho to a transcript to be terminated

For Rho to terminate the synthesis of a transcript, it first

must become attached in a productive manner. A segment of

the nascent transcript must be available for Rho to bind with

a sufficiently high affinity. In bacteria, ribosomes become

attached to a nascent mRNA as soon the translation initia-

tion site is available and this is usually before Rho can

attach to the RNA. The presence of a ribosome translating

the mRNA at the same rate with which the RNA is being

synthesized effectively prevents Rho from access to the

nascent RNA until the end of the gene or the operon [35].

J.P. Richardson / Biochimica et Biophysica Acta 1577 (2002) 251–260 253

Once the coupling is interrupted, the nascent RNA is then

available, and, if the sequence is appropriate, Rho will

become attached and initiate the termination process. An

appropriate sequence consists of a segment of 40 or more

non-base-paired nucleotides containing a relatively high

proportion of C residues [36,37]. Such a sequence is called

a rut site, named after the genetic designation of the DNA

sequence element called the Rho Utilization site [38]. This

site on the RNA has also been referred to as the rut site for

the RNA attachment role it actually performs [39].

One example of a Rho-dependent terminator is tR1, the

transcript terminator between the cro and cII genes of

bacteriophage E [40]. Its rut sequence starts just down-

stream from the open reading frame for the Cro protein and

just upstream of where transcripts are terminated by Rho

action [38,41]. Rho binds to the E cro RNA containing its

rut sequence with Kdf 30 pM [42]. In contrast the affinity

of Rho to E cro RNAvariants lacking the rut site segment or

to the RNA containing the rut sequence but with it base-

paired to complementary DNA oligonucleotides is about

100-fold weaker [43]. These results correlate with the

findings that deletion of the rut segment of DNA or replace-

ment of the segment of the DNA with one that forms a

highly base-paired structure lead to a loss of termination

function both in vitro and in vivo [38,44]. In addition,

blocking the access to the rut site in vitro by the addition

of DNA oligonucleotides that are complementary specifi-

cally to the rut sequence segments on the nascent RNA also

strongly inhibits Rho-dependent termination [41].

5. Termination sites

A Rho-dependent terminator consists of two parts, the rut

region and the region where transcripts are terminated,

called the tsp region [45]. This second part can extend for

nearly 100 base pairs, thus giving an extent of nearly 150 bp

overall for a Rho-dependent terminator. The broad extent of

the tsp region is because the transcripts are terminated at

several positions often in clusters of five to eight successive

stop points separated by regions of 20–30 bp where few

transcripts are terminated [40,46,47]. This dispersion of the

stop points differs from the relatively precise position of the

stop points for an intrinsic terminator, in which most tran-

scripts end at a single base pair with a small fraction at an

adjacent base pair [2].

The positions of the stop points in a Rho-dependent

terminator are determined by the sequence features that

control the rate of nucleotide addition by RNA polymerase

at each residue. The rate for addition of nucleotides by RNA

polymerases can vary considerably from one position to the

other, giving rise to characteristic pause points that are

distributed very much like the stop points at a Rho-depend-

ent terminator [48]. These pause points are dictated by

several features of the DNA sequence in and around the

base pairs at which the pausing occurs [49,50]. Comparing

the distribution of natural pause points in a tsp region of a

Rho-dependent terminator during transcription of the region

in the absence of Rho with the distribution of stop points of

the RNA molecules synthesized in the presence of Rho

shows a very good correlation [51,52]. Thus, termination

occurs at pause sites that are downstream from a rut region.

Pause sites occur naturally and with similar frequencies

in most DNA sequences. As a consequence, when a new

DNA sequence not from a natural termination site is

positioned downstream from a rut region, Rho will cause

transcripts to be terminated at the natural pause sites in the

inserted DNA [45]. This characteristic implies that the most

unique determinant of a Rho-dependent terminator is the rut

region. Also, because Rho does not bind stably to an RNA

until most of the rut region is available [53], it acts very

efficiently at the first pause sites encountered by RNA

polymerase just after the rut region is available. This means

that the distance between where Rho loads on the RNA and

where the transcript is terminated is relatively short, in a

range from 20 to 40 nucleotides. Models showing Rho

binding to a far upstream site and having it migrate along

scores of RNA nucleotides until it encounters a paused RNA

polymerase are probably not good representations.

Mutant forms of Rho that act more slowly than normal

Rho are very poor at terminating transcription with a normal

RNA polymerase under normal transcription conditions

[54]. However, the function of these mutants can be recov-

ered when they are used with a mutant RNA polymerase

that elongates RNA chains more slowly or with a normal

RNA polymerase under conditions of lower NTP concen-

trations [55]. These observations reveal a kinetic coupling

between Rho action and transcript elongation. A critical

aspect of this kinetic coupling is the timing of how long it

takes Rho to start tracking along a transcript once it has

bound tightly to the transcript. Even if the distance to the

pause site is not far, a delay in acting on the RNAwill cause

Rho to lose its chance to dissociate the transcript before

RNA polymerase has moved beyond the pause site.

6. ATP binding and hydrolysis

Rho–RNA complexes catalyze the hydrolysis of all four

nucleoside triphosphates [56]. However, its preferred sub-

strate is ATP. Rho will bind to ATP in the absence of RNA.

Each hexamer has three sites that bind ATP with Kdf 0.5

AM [57]. Although there was some evidence that Rho could

bind three more ATP molecules with a much weaker affinity

[58,59], the existence of these sites has been difficult to

verify [60]. Pre-steady state kinetic measurements using a

rapid mix and quench device showed that addition of RNA

(poly(C)) to the Rho–ATP3 complex led to hydrolysis of 1

ATP in less than 5 ms and at least 1.5 more ATP by 200 ms

[60]. These results and others measuring the rates of

hydrolysis of labeled ATP added to a presaturated Rho–

ATP3 complex have been simulated by a model with the


following properties [60]. Rho has three catalytic sites in

which ATP can bind. Addition of RNA to the Rho–ATP3complex causes rapid hydrolysis (rate constant of 300 s� 1)

of one ATP. This reaction is followed by a rate-limiting step

(either product release or protein conformational change) at

27 s � 1. Additional bound ATP molecules sequentially

undergo similar rapid chemical hydrolysis followed by a

similar slow step. This model is consistent with the steady-

state hydrolysis rate (turnover number) of 30 s � 1 per

hexamer. The rapid quench results were not consistent with

an alternative model that has been proposed in which ATP

bound at slow exchanging sites (i.e. the high affinity sites) is

hydrolyzed at a relatively slow rate of 1.8 s� 1 [14]. That

alternative model posits that the high-affinity sites are not

catalytic and that turnover ATP hydrolysis is carried out in

the putative low-affinity sites. Attempts to reproduce the

experiments that were consistent with this alternative model

in another laboratory were not successful [60], suggesting

that those results may have been a characteristic of a

particular preparation of Rho protein.

7. The role of NTP hydrolysis

As noted above, Rho can bind to RNA in the absence of

an NTP. However, in the cell and in transcription assay

mixtures, NTPs are present and used by Rho as substrates to

mediate the termination process. The interaction of Rho

with RNA activates catalytic hydrolysis of NTPs to the

corresponding NDP and inorganic phosphate (Pi). When the

RNA is a nascent transcript, the hydrolysis reaction

becomes coupled to the release of the transcript and hence

its termination [8,9]. However, even after the transcript has

been released, Rho will continue to interact with it and to

hydrolyze ATP at a steady-state turnover rate of as much as

60 NTPs s� 1 per Rho [29,45]. The hydrolysis reaction is

absolutely dependent on RNA. Even though some single-

stranded DNA molecules can bind to Rho nearly as tightly

as RNA can, those DNA molecules do not activate any ATP

hydrolysis [24]. On the other hand, the release reaction is

not limited to polymerase and DNA template attached to the

3V end of a transcript. Rho will also catalyze the release of

DNA molecules or DNA oligonucleotides that are paired

with a segment (20–40 base pairs) of the RNA, as long as

the DNA is on the 3V side of the rut segment of the RNA

[61,62]. This release is again coupled to NTP hydrolysis.

Thus, this ability to act as a hybrid helicase indicates that

Rho can move on a transcript in a direction that will bring it

into contact with the RNA polymerase that is synthesizing

the RNA. This tracking action on the RNA could also be the

driving force that pulls the transcript from its complex with

the polymerase and the DNA template.

Currently, there is a dispute over the nature of the

tracking reaction. In one interpretation, which is supported

by an elegant experiment [63], Rho maintains a contact with

the rut site used for its initial tight binding to the transcript

[64]. In this mechanism, Rho remains tethered to the rut site

on the RNA while it is tracking toward the 3V end of that

same RNA [63]. In an alternative view [65], Rho is

proposed to migrate down the RNA without maintaining

its original contact. Because it is difficult to prove that one

Rho is making separate contacts at two distinct regions of an

RNA during the migration process, neither mechanism can

be eliminated at this point.

Part of the dispute is on the number and placement of

sites on Rho for binding to RNA. Rho appears to have two

kinds of sites for binding RNA. This became evident with

the finding that ATP hydrolysis by Rho could be activated

by combinations of single-stranded DNA and short oligor-

ibonucleotides [66]. One kind of site binds both RNA and

DNA while the other site is specific for RNA. NTP

hydrolysis would then be dependent upon interactions at

both sites. It is well established that both RNA and DNA

can bind to the OB-fold segment of the RNA-BD, thus

placing the primary site in that domain [33]. However, until

recently, the location of the secondary site was not known.

One reasonable suggestion was that the two kinds of sites

could be different conformations of the site, the OB fold, in

the RNA-BD in different subunits of hexameric Rho [65].

This interpretation is supported by the finding that three of

the subunits can bind oligo(C) with high affinity and three

bind it with lower affinity [31]. However, two lines of recent

evidence suggest that RNA can also interact with a site that

is in the ATP-BD.

The first line of evidence is from cross-linking studies, in

which a photoactivatable chemical group on a loop region

(the R-loop, see Fig. 1) of the ATP-BD form cross-links to

RNA if the RNA has a rut sequence and an RNA segment

on the 3V of the rut sequence [23]. In contrast, an RNA that

has its 3V end at the end of the rut sequence forms cross-

links only with a group placed in the OB fold of the RNA-

BD. The position of the R loop in the subunits of the

homology-modeled structure of Rho is in the hole in the

center of the ring-shaped structure. This result suggests that

the rut segment of the RNA makes contacts in the RNA-BD

while RNA to the 3V of rut passes through the hole of the

Rho hexamer. This passage of a polynucleotide segment

through the hole of a ring-shaped hexamer is a characteristic

of hexameric DNA helicases, proteins that are structurally

and functionally similar to Rho [67–69].

The second line of evidence has come from studies of the

protection of Rho from cleavage by reactive peroxyl-species

generated by the interaction of chelated iron with hydrogen

peroxide [70]. An exposed part of Rho on a loop in the ATP-

BD, in this case the Q-loop (see Fig. 1), which also faces

into the hole of the Rho hexamer, becomes protected

specifically by RNA, not DNA. On the other hand, both

RNA and DNA gave extensive protection of accessible parts

of the OB-fold in the RNA-BD. These results suggest the

presence of an RNA-specific site on or near the Q-loop. The

amino acid residues on this loop are well conserved in the

Rho homologues, and mutational studies have now demon-


strated that changes of any of the four Q-loop-residues that

can make hydrogen bonds or ionic interactions to alanines

abrogate Rho termination function [70a]. More specifically,

these mutants are very defective in their interactions with

RNA in their secondary site but not in their primary site.

Taken together, the results of the protection studies and the

functional analysis of the Q-loop mutations indicate that the

Q-loop plays an important role in the interactions of Rho

with RNA. They are also consistent with the model that has

the 3V segment of RNA passing through the hole in the ring-

shaped Rho hexamer. Fig. 2 is a diagram showing the path

of an RNA molecule bound to a Rho hexamer, showing the

extensive contact of a portion of the RNAwith the extended

cleft formed by the RNA-BD and the 3Vsegment passing

through the hole in the center. The structure of Rho used

here is based on the EM image reconstruction.

8. Model for termination of a transcript with Rho

Based on the mechanism that has been proposed for the

tracking of Rho on an RNA molecule, I propose the

following model for termination of a transcript with Rho.

As soon as a sufficient amount of a rut segment of RNA has

emerged from the exit site of RNA polymerase, Rho binds

to the RNA by making contacts with the OB folds in several

domains of the hexameric or mainly hexameric Rho. This

initial interaction would include high-affinity contacts in at

least two of the subunits with lower affinity contacts with

two other subunits. Even when bound to RNA, a subunit

can dissociate, allowing the segment of the RNA on the 3Vside of rut to be captured in the opening to the center of the

ring. Rebinding of that subunit would close the ring and set

the stage for ATP hydrolysis. The rut segment of the

transcript would be bound in a cleft that extends around

the periphery of one side of the hexamer, the side facing

away from RNA polymerase (Fig. 3). The part of Rho in

contact with the RNA polymerase would be the other face

and would consist of the six C-terminal portions of the ATP-

BD ringing the exit hole.

The simultaneous contact of an RNAwith OB folds in the

RNA-BD and with the Q-loop on one of the six subunits in

the narrowest part of the hole in the ring would then trigger

ATP hydrolysis in that subunit. The conversion to the ADP

form or the subsequent release of the ADP and Pi would be

coupled to a conformational change in that subunit. This

change could pass the interaction of the RNA in the hole to

the Q-loop of the next subunit containing an ATP molecule.

This would go in one specific direction. After hydrolysis of

that ATP, the RNA segment would be passed in that same

direction to the third subunit of the three that bind ATP.

Coupled with this sequential and rotational hydrolysis of

ATP molecules will be an overall conformation shift that is

coupled to the translocation of the RNA in the 5V to 3Vdirection through the hole. One possible way for the trans-

location to occur is by a screwing action on the RNA in

which the hole of the Rho hexamer is acting conformation-

Fig. 2. Topological model of Rho bound to an mRNA. The shape of the

hexameric Rho is based on the three-dimensional reconstruction of electron

micrograph images as reported by Yu et al. [10]. The rut segment of the

RNA is bound in the continuous cleft that extends around the upper

periphery. The 3Vsegment of the RNA passes through the center ending in

the active site of the RNA polymerase (not shown).

Fig. 3. Diagram of a transcription elongation complex (TEC) based on the model of Korzheva et al. [103] before and after attachment of Rho to the nascent

RNA. After attachment, Rho acts to pull the nascent transcript from the elongation complex. The 5V segment of the RNA (brown line) is shown wrapped

around the binding site at one end of the orange-shaped Rho structure (shown in orange). Note that the RNA has lost some of its pairing with the template

(green line) and is thus just about to be pulled away by Rho. The arrows on the left image indicate the direction of translocation of the DNA template.


ally like a nut on a helical single-stranded RNA. This action

could pull the RNA through with a step of about one

nucleotide per ATP hydrolyzed, as has been measured

[71], or at a linear translocation rate as fast as 60 nucleotides

per second (based on the ATP hydrolysis turnover number).

This estimated rate is in the right range to be a bit faster than

the average rate of transcript elongation of about 50 nucleo-

tides per second [1]. Hence, when RNA polymerase reaches

a pause site after Rho is firmly attached, the rate would be

fast enough to pull the RNA out of the RNA polymerase.

The dissociation of the RNA from its base-pairing inter-

actions with the template DNA could be by one of the

proposed mechanisms for helicases [72]. The bases paired

at the ends of double helices are known to separate briefly as

a result of thermal motions (called breathing). If Rho pulls

the RNA into its hole by a nucleotide step during the

temporary base pair separation, it will capture the RNA in

a form that will not be able to reform that terminal pair with

the DNA template. This could happen as Rho is pulling at the

RNA at the exit site with the contact (perhaps nonspecific) of

Rho with RNA polymerase acting as the leverage for the

pulling action. Step by step the pairing with the template is

broken and the RNA pulled completely through the exit site

to affect the release of the transcript. Fig. 3 shows a diagram

of a Rho molecule pulling the transcript partially away from

its interactions with the DNA template in RNA polymerase.

9. NusG is a Rho cofactor

In the cell, the function of Rho at some terminators

depends on the presence of a 21 kDa monomeric protein

called NusG [73]. This protein was discovered because of its

function as a host cell cofactor for the transcription anti-

termination function of bacteriophage E protein N [74].

However, it has since been found to be a transcript elonga-

tion factor [75], which acts to enhance the escape of RNA

polymerase from class II pause sites [49] and has the general

effect of increasing the rate of transcript elongation by about

25% [76]. NusG is an essential protein for viability of E.

coli [77]. In cells that are experimentally depleted of NusG,

some Rho-dependent terminators no longer function prop-

erly, though others seem less affected [73]. A requirement of

NusG for Rho function was overlooked, presumably

because the conditions that were used to study Rho function

in vitro had been optimized for Rho function by reduction of

the NTP concentration. When transcripts are elongated in

vitro at the same rate that occurs in vivo, some terminators

do show a dependence on NusG for efficient Rho-mediated

termination [78]. These results suggest that NusG is over-

coming a kinetic deficiency of Rho to act by itself under

certain conditions.

NusG binds both RNA polymerase [79] and Rho [26],

although it is not known yet whether it can bind both

simultaneously. Assuming that it can, it could act either to

bring Rho into close proximity to the emerging transcript or

to stabilize the complex of Rho with the nascent transcript

and RNA polymerase [80]. By forming a complex of Rho

with RNA polymerase before the rut site emerges, the

binding of the rut segment to the primary RNA-binding site

would become a unimolecular reaction. Thus, this hypoth-

esis is consistent with the evidence that NusG is acting to

overcome a kinetic deficiency of Rho to act by itself.

NusG has been shown to make a 1:1 complex with

hexameric Rho [27]. The position of this unique binding

site on the Rho hexamer has not been located, but because it

is unique, it is likely to be in one of the two entrances to the

hole. If a NusG bound to RNA polymerase also binds to

Rho on the side with the RNA-BD, it would orient Rho so

that the emerging transcript would be in close proximity to

the cleft of the RNA-BD. However, once the transcript is

bound tightly in part of that cleft, that interaction with NusG

would have to be released to allow the RNA to enter the

hole. In addition Rho would have to rotate 180j end to end

to have the hole at the C-terminal domains facing RNA

polymerase as is shown in the diagram in Fig. 3. This would

be some fancy footwork. An alternative suggested role for

NusG is that it favors an RNA polymerase conformation

that enhances RNA release [27,81].

NusA protein has been shown to bind to Rho and RNA

polymerase and can do so simultaneously [28]. NusA, like

NusG, is a host factor for bacteriophage E protein N-

mediated anti-termination. It is also a transcript elongation

factor, but unlike NusG, NusA acts to enhance pausing at a

certain class of pause sites and thus slows the rate of

transcript elongation [49,51]. Its action in Rho-dependent

termination has been ambiguous. In most assays, it acts to

inhibit Rho function, perhaps by competing with Rho for rut

segment interactions [40,76,82]. However, there is also a

report in which NusA is shown to enhance Rho action [83],

as might be expected from its ability to enhance pausing.

NusA is essential for viability of E. coli [84]. However, a

conditionally lethal mutation in nusA can be suppressed by

certain mutations in rho, namely mutations that allow

expression of an active but defective Rho protein [85]. This

observation suggests that the functions of the two proteins

are competitive. One possible role of NusA is to ensure a

close coupling of translation with transcription, perhaps by

retarding transcript elongation to prevent RNA polymerase

from moving ahead of a translating ribosome. Although this

is an attractive model and is consistent with the suppression

of a NusA defect by a Rho defect, direct evidence for this

function is lacking.

10. Biological role of Rho

As mentioned in the introduction, E. coli uses two kinds

of transcript terminators. This is not true for all bacteria.

With the advent of genome sequencing, it is now known that

a few bacteria, a Mycoplasma and a Synecchocystis for

instance, lack a rho gene. On the other hand, most bacteria


have a rho gene. It is present in organisms in lineages that

diverged from that of E. coli over a billion years ago,

indicating that it is not a recent addition [21]. However,

there is no evidence yet for a rho-like gene in archael or

eukaroytic organisms. Thus, Rho is not essential for life or

even life in the bacteria, but it arose early in evolution,

suggesting that it has a useful function for most bacteria.

The question of whether eukaroytic cells use an ATP-

dependent mechanism for termination of transcripts is

considered in the final section of this review.

The rho gene is essential for E. coli and a few other

organisms [86,87]. In general, cells that are sensitive to the

Rho-specific antibiotic bicyclomycin probably have essen-

tial rho genes [88,89]. However, rho is not essential for

Bacillus subtilis [90] or for Staphylococcus aureus [91]. In

B. subtilis, rho is expressed but the level of Rho in the cells

is 50-fold lower than the level of Rho in E. coli [92]. The

Rho that is present does function at least to regulate its own

synthesis. Altogether, these observations suggest that B.

subtilis does not make extensive use of Rho-dependent

terminators for the orderly expression of its genes, but

nonetheless maintains the gene and expresses it for some

function.

The extent to which E. coli makes use of its two kinds

of transcript terminators is not known. Putative intrinsic

terminators can be identified by genome analysis, but the

actual efficiency of functioning of a sequence as a termi-

nator is still difficult to predict. For example, a sequence

for an intrinsic terminator is found at the end of the trp

operon of E. coli [93]. However, functional studies sub-

sequently showed that that terminator was very weak and

that a much more efficient Rho-dependent terminator was

responsible for terminating most transcripts for that operon

[47]. A similar organization was found in a number of other

operons [94]. Because Rho-dependent terminators lack a

readily identifiable consensus sequence, it is not known

how frequently they are used. Depletion of a cell of Rho

may be lethal because of the catastrophic effect of the

expression of many different sequences that normally are

not expressed or because products of the inadvertent

expression are toxic.

The fact that the rho gene arose early in the natural

history of the bacteria suggests that it gives some selective

advantage to a cell. One likely advantage is that under

conditions of stress, it prevents the continued synthesis of

an RNA that cannot be translated [35]. In E. coli, Rho has

been shown to terminate the synthesis of transcripts very

early in the lac operon when the cells are starved for amino

acids [95]. This comes about because the lac operon, like

many other operons, contains latent Rho-dependent termi-

nators within the early part of the gene [96]. These intra-

genic terminators do not function under conditions of

normal expression, presumably because a ribosome is

present translating the RNA as it emerges from the exit

site of RNA polymerase. This ribosome would block the

access of Rho to the RNA in a way that would allow it to

terminate transcription. However, when the movement of

the ribosome is slowed or blocked because of the absence

of an amino acid, a segment of a transcript containing a rut

site would become exposed allowing Rho to bind and

terminate the partial transcript by its normal mechanism.

The selective advantage this would have is that it would

prevent the loss of energy in making a transcript that will

not be translated.

The broad sequence specificity of a Rho dependent

terminator as well as the dependence of access of Rho to

the transcript makes this type of terminator well suited for

this stress-protection function. These features also made

Rho-dependent terminators very suitable for placement at

the end of a gene or operon.

11. Is a Rho factor present in eukaryotic cells?

A factor that is analogous to Rho has not yet been

discovered in eukaryotic cells. These cells certainly lack a

gene that has the strongly conserved sequence features that

are characteristic of the rho genes in bacteria. However,

one aspect of transcript termination with RNA polymerase

II is similar to the Rho-dependent mechanism and that is

termination occurs in broad regions downstream from a

sequence signal [97,98]. Thus, these cells could have

another kind of protein with the ability to bind to a

transcript and act as a molecular machine to translocate

along the RNA and wrest the transcript from RNA poly-

merase. An ATPase, called factor 2, which acts to disso-

ciate transcripts from RNA polymerase II, has been isolated

[99], but its function may be involved with release of

abortive products at the early stage of the transcription

process. It is not an RNA-binding protein. Instead, its

ATPase activity is stimulated by DNA [100]. Also, its

sequence makes it related to chromatin remodeling proteins

[101]. RNA polymerase I also makes use of a protein

factor, called Reb1p in yeast, to mediate release of tran-

scripts at terminators, but this factor binds to DNA and

does not appear to require ATP hydrolysis [102].

Acknowledgements

I thank Ronnie Wei for preparation of Figs. 1 and 2 and

Lislott Richardson, R. Wei and Brandt Burgess for

discussions. Research cited from my laboratory was

supported by NIH Grant GM56095.

References

[1] J.P. Richardson, J. Greenblatt, in: F.C. Neidhardt, R. Curtiss III, J.L.

Ingraham, E.C.C. Lin, K.B. Low, B. Magasanik, W.S. Reznikoff, M.

Riley, M. Schaechter, H.E. Umbarger (Eds.), Escherichia coli and

Salmonella: Cellular and Molecular Biology, 2nd Ed., Washington,

DC, 1996, pp. 822–848.


[2] K.S. Wilson, P.H. von Hippel, J. Mol. Biol. 244 (1994) 36–51.

[3] R.A. Mooney, I. Artsimovitch, R. Landick, J. Bacteriol. 180 (1998)

3265–3275.

[4] I. Gusarov, E. Nudler, Mol. Cell 3 (1999) 495–504.

[5] W.S. Yarnell, J.W. Roberts, Science 284 (1999) 611–615.

[6] Y. dVAubenton Carafa, E. Brody, C. Thermes, J. Mol. Biol. 216

(1990) 835–858.

[7] F.H. Martin , I. Tinoco Jr., Nucleic Acids Res. 8 (1980) 2295–2299.

[8] K. Shigesada, C.W. Wu, Nucleic Acids Res. 8 (1980) 3355–3369.

[9] J.P. Richardson, R. Conaway, Biochemistry 19 (1980) 4293–4299.

[10] X. Yu, T. Horiguchi, K. Shigesada, E.H. Egelman, J. Mol. Biol. 299

(2000) 1279–1287.

[11] L.R. Finger, J.P. Richardson, J. Mol. Biol. 156 (1982) 203–219.

[12] J. Geiselmann, T.D. Yager, S.C. Gill, P. Calmettes, P.H. von Hippel,

Biochemistry 31 (1992) 111–121.

[13] B.R. Burgess, J.P. Richardson, J. Biol. Chem. 276 (2001)

17106–17110.

[14] D.E. Kim, S.S. Patel, J. Biol. Chem. 274 (1999) 32667–32671.

[15] J.L. Pinkham, T. Platt, Nucleic Acids Res. 11 (1983) 3531–3545.

[16] D. Modrak, J.P. Richardson, Biochemistry 33 (1994) 8292–8299.

[17] T.J. Allison, T.C. Wood, D.M. Briercheck, F. Rastinejad, J.P.

Richardson, G.S. Rule, Nat. Struct. Biol. 5 (1998) 352–356.

[18] A.J. Dombroski, J.R. LaDine, R.L. Cross, T. Platt, J. Biol. Chem.

263 (1988) 18810–18815.

[19] J.W. Dolan, N.F. Marshall, J.P. Richardson, J. Biol. Chem. 265

(1990) 5747–5754.

[20] D.M. Briercheck, T.C. Wood, T.J. Allison, J.P. Richardson, G.S.

Rule, Nat. Struct. Biol. 5 (1998) 393–399.

[21] T. Opperman, J.P. Richardson, J. Bacteriol. 176 (1994) 5033–5043.

[22] Y. Miwa, T. Horiguchi, K. Shigesada, J. Mol. Biol. 254 (1995)

815–837.

[23] B.R. Burgess, J.P. Richardson, J. Biol. Chem. 276 (2000) 4182–4189.

[24] G.R. Galluppi, J.P. Richardson, J. Mol. Biol. 138 (1980) 513–539.

[25] J.A. McSwiggen, D.G. Bear, P.H. von Hippel, J. Mol. Biol. 199

(1988) 609–622.

[26] J. Li, S.W. Mason, J. Greenblatt, Genes Dev. 7 (1993) 161–172.

[27] Z. Pasman, P.H. von Hippel, Biochemistry 39 (2000) 5573–5585.

[28] M.C. Schmidt, M.J. Chamberlin, J. Biol. Chem. 259 (1984)

15000–15002.

[29] C. Lowery-Goldhammer, J.P. Richardson, Proc. Natl. Acad. Sci.

U. S. A. 71 (1974) 2003–2007.

[30] D.G. Bear, P.S. Hicks, K.W. Escudero, C.L. Andrews, J.A.

McSwiggen, P.H. von Hippel, J. Mol. Biol. 199 (1988) 623–635.

[31] J. Geiselmann, T.D. Yager, P.H. von Hippel, Protein Sci. 1 (1992)

861–873.

[32] Y. Wang, P.H. von Hippel, J. Biol. Chem. 268 (1993) 13947–13955.

[33] D.M. Briercheck, T.J. Allison, J.P. Richardson, J.F. Ellena, T.C.

Wood, G.S. Rule, J. Biomol. NMR 8 (1996) 429–444.

[34] C.E. Bogden, D. Fass, N. Bergman, M.D. Nichols, J.M. Berger, Mol.

Cell 3 (1999) 487–493.

[35] J.P. Richardson, Cell 64 (1991) 1047–1049.

[36] P. Alifano, F. Rivellini, D. Limauro, C.B. Bruni, M.S. Carlomagno,

Cell 64 (1991) 553–563.

[37] W.D. Morgan, D.G. Bear, B.L. Litchman, P.H. von Hippel, Nucleic

Acids Res. 13 (1985) 3739–3754.

[38] C.Y. Chen, J.P. Richardson, J. Biol. Chem. 262 (1987) 11292–11299.

[39] D.G. Bear, D.S. Peabody, Trends Biochem. Sci. 13 (1988) 343–347.

[40] L.F. Lau, J.W. Roberts, R. Wu, Proc. Natl. Acad. Sci. U. S. A. 79

(1981) 6171–6175.

[41] C.Y. Chen, G.R. Galluppi, J.P. Richardson, Cell 46 (1986) 1023–1028.

[42] E. Gan, J.P. Richardson, Biochemistry 38 (1999) 16882–16888.

[43] I. Faus, J.P. Richardson, Biochemistry 28 (1989) 3510–3517.

[44] J.E. Graham, J.P. Richardson, J. Biol. Chem. 273 (1998) 20764–20769.

[45] L.V. Richardson, J.P. Richardson, J. Biol. Chem. 271 (1996)

21597–21603.

[46] W.D. Morgan, D.G. Bear, P.H. von Hippel, J. Biol. Chem. 258

(1983) 9553–9564.

[47] A.M. Wu, G.E. Christie, T. Platt, Proc. Natl. Acad. Sci. U. S. A. 78

(1981) 2913–2917.

[48] J.R. Levin, M.J. Chamberlin, J. Mol. Biol. 196 (1987) 61–84.

[49] I. Artsimovitch, R. Landick, Proc. Natl. Acad. Sci. U. S. A. 97

(2000) 7090–7095.

[50] C.L. Chan, R. Landick, J. Mol. Biol. 233 (1993) 25–42.

[51] L.F. Lau, J.W. Roberts, R. Wu, J. Biol. Chem. 258 (1983) 9391–9397.

[52] W.D. Morgan, D.G. Bear, P.H. von Hippel, J. Biol. Chem. 258

(1983) 9565–9574.

[53] M.A. Ceruzzi, S.L. Bektesh, J.P. Richardson, J. Biol. Chem. 260

(1985) 9412–9418.

[54] J.P. Richardson, J.L. Carey, J. Biol. Chem. 257 (1982) 5767–5771.

[55] D.J. Jin, R.R. Burgess, J.P. Richardson, C.A. Gross, Proc. Natl.

Acad. Sci. U. S. A. 89 (1992) 1453–1457.

[56] C. Lowery, J.P. Richardson, J. Biol. Chem. 252 (1977) 1375–1380.

[57] B.L. Stitt, J. Biol. Chem. 263 (1988) 11130–11137.

[58] J. Geiselmann, P.H. von Hippel, Protein Sci. 1 (1992) 850–860.

[59] D.E. Kim, K. Shigesada, S.S. Patel, J. Biol. Chem. 274 (1999)

11623–11628.

[60] B.L. Stitt, Biochemistry 40 (2001) 2276–2281.

[61] C.A. Brennan, A.J. Dombroski, T. Platt, Cell 48 (1987) 945–952.

[62] K.M. Walstrom, J.M. Dozono, S. Robic, P.H. von Hippel, Biochem-

istry 36 (1997) 7980–7992.

[63] E.J. Steinmetz, T. Platt, Proc. Natl. Acad. Sci. U. S. A. 91 (1994)

1401–1405.

[64] I. Faus, J.P. Richardson, J. Mol. Biol. 212 (1990) 53–66.

[65] J. Geiselmann, Y. Wang, S.E. Seifried, P.H. von Hippel, Proc. Natl.

Acad. Sci. U. S. A. 90 (1993) 7754–7758.

[66] J.P. Richardson, J. Biol. Chem. 257 (1982) 5760–5766.

[67] M.J. Jezewska, S. Rajendran, W. Bujalowski, Biochemistry 37

(1998) 3116–3136.

[68] H.H. Egelman, X. Yu, R. Wild, M.M. Hingorani, S.S. Patel, Proc.

Natl. Acad. Sci. U. S. A. 92 (1995) 3869–3873.

[69] S.S. Patel, K.M. Picha, Annu. Rev. Biochem. 69 (2000) 651–697.

[70] R.R. Wei, J.P. Richardson, J. Biol. Chem. 276 (2001) 28380–28387.

[71] R.R. Wei, J.P. Richardson, J. Mol. Biol. 314 (2001) 1007–1015.

[72] K.M. Walstrom, J.M. Dozono, P.H. von Hippel, Biochemistry 36

(1997) 7993–8004.

[73] P.H. von Hippel, E. Delagoutte, Cell 104 (2001) 177–190.

[74] S.L. Sullivan, M.E. Gottesman, Cell 68 (1992) 989–994.

[75] S.W. Mason, J. Greenblatt, Genes Dev. 5 (1991) 1504–1512.

[76] E. Burova, S.C. Hung, V. Sagitov, B.L. Stitt, M.E. Gottesman, J.

Bacteriol. 177 (1995) 1388–1392.

[77] C.M. Burns, L.V. Richardson, J.P. Richardson, J. Mol. Biol. 278

(1998) 307–316.

[78] W.L. Downing, S.L. Sullivan, M.E. Gottesman, P.P. Dennis, J. Bac-

teriol. 172 (1990) 1621–1627.

[79] C.M. Burns, J.P. Richardson, Proc. Natl. Acad. Sci. U. S. A. 92

(1995) 4738–4742.

[80] J. Li, R. Horwitz, S. McCracken, J. Greenblatt, J. Biol. Chem. 267

(1992) 6012–6019.

[81] K.W. Nehrke, T. Platt, J. Mol. Biol. 243 (1994) 830–839.

[82] C.M. Burns, W.L. Nowatzke, J.P. Richardson, J. Biol. Chem. 274

(1999) 5245–5251.

[83] C.D. Sigmund, E.A. Morgan, Biochemistry 27 (1988) 5622–5627.

[84] M. Kainz, R.L. Gourse, J. Mol. Biol. 284 (1998) 1379–1390.

[85] M.G. Craven, D.I. Friedman, J. Bacteriol. 173 (1991) 1485–1491.

[86] C. Zheng, D.I. Friedman, Proc. Natl. Acad. Sci. U. S. A. 91 (1994)

7543–7547.

[87] S. Brown, E.R. Brickman, J. Beckwith, J. Bacteriol. 146 (1981)

422–425.

[88] A. Das, D. Court, S. Adhya, Proc. Natl. Acad. Sci. U. S. A. 73

(1976) 1959–1963.

[89] A. Zwiefka, H. Kohn, W.R. Widger, Biochemistry 32 (1993)

3564–3570.

[90] W.L. Nowatzke, E. Keller, G. Koch, J.P. Richardson, J. Bacteriol. 179

(1997) 5238–5240.


[91] P.G. Quirk , E.A. Dunkley Jr., P. Lee, T.A. Krulwich, J. Bacteriol. 175

(1993) 647–654.

[92] R.S. Washburn, A. Marra, A.P. Bryant, M. Rosenberg, D.R. Gentry,

Antimicrob. Agents Chemother. 45 (2001) 1099–1103.

[93] C.J. Ingham, J. Dennis, P.A. Furneaux, Mol. Microbiol. 31 (1999)

651–663.

[94] A.M.Wu, T. Platt, Proc.Natl. Acad. Sci. U. S.A. 75 (1978) 5442–5446.

[95] J.H. Sameshima, R.C. Wek, G.W. Hatfield, J. Biol. Chem. 264

(1989) 1224–1231.

[96] E.C. Ruteshouser, J.P. Richardson, J. Mol. Biol. 208 (1989) 23–43.

[97] S. Adhya, M. Gottesman, B. de Crombrugghe, D. Court, in: M.

Chamberlin, R. Losick (Eds.), Transcription Termination Regulates

Gene Expression, Cold SpringHarbor, NewYork, 1976, pp. 719–730.

[98] N.J. Proudfoot, Trends Biochem. Sci. 14 (1989) 105–110.

[99] S. Connelly, J.L. Manley, Genes Dev. 2 (1988) 440–452.

[100] Z. Xie, D.H. Price, J. Biol. Chem. 271 (1996) 11043–11046.

[101] Z. Xie, D.H. Price, J. Biol. Chem. 273 (1998) 3771–3777.

[102] M. Liu, Z. Xie, D.H. Price, J. Biol. Chem. 273 (1998) 25541–25544.

[103] W.H. Lang, R.H. Reeder, Proc. Natl. Acad. Sci. U. S. A. 92 (1995)

9781–9785.

[104] N. Korzheva, A. Mustaev, M. Kozlov, A. Malhotra, V. Nikiforov, A.

Goldfarb, S.A. Darst, Science 289 (2000) 619–625.


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Rho-dependent termination and ATPases in transcript termination