DOI 10.1515/hsz-2013-0133      Biol. Chem. 2013; 394(7): 845–855

Review

Ü lo Maiv ä li * , Anton Paier and Tanel Tenson

When stable RNA becomes unstable: the degradation of ribosomes in bacteria and beyond Abstract: This review takes a comparative look at the

various scenarios where ribosomes are degraded in bac-

teria and eukaryotes with emphasis on studies involving

Escherichia coli and Saccharomyces cerevisiae . While the

molecular mechanisms of degradation in bacteria and

yeast appear somewhat different, we argue that the under-

lying causes of ribosome degradation are remarkably sim-

ilar. In both model organisms during ribosomal assembly,

partially formed pre-ribosomal particles can be degraded

by at least two different sequentially-acting quality con-

trol pathways and fully assembled but functionally faulty

ribosomes can be degraded in a separate quality control

pathway. In addition, ribosomes that are both structur-

ally- and functionally-sound can be degraded as an adap-

tive measure to stress.

Keywords: degradation; Escherichia coli ; nonfunctional

RNA decay (NRD); ribosome; yeast.

*Corresponding author: Ü lo Maiv ä li, Institute of Technology,

University of Tartu, Nooruse 1, Tartu 50411, Estonia,

e-mail: ymaivali@ut.ee

Anton Paier and Tanel Tenson: Institute of Technology, University of

Tartu, Nooruse 1, Tartu 50411, Estonia

Introduction Traditionally, cellular RNAs have been described as either

unstable or stable. Messenger RNAs (mRNAs) are typical

of unstable RNAs, with lifetimes from minutes to hours,

while ribosomal RNAs (rRNAs) and transfer RNAs (tRNAs)

are often viewed as stable, with lifetimes that can exceed

several weeks (Metodiev et  al. , 2009 ). This dichotomy

of stability regimes is intuitively well supported by the

notion that it must be advantageous to maintain a rapid

turnover of mRNA to enable rapid suppression of the syn-

thesis of individual proteins by stopping transcription of

their genes. In contrast, RNAs that perform housekeeping

functions should ideally live forever to reduce the cost of

making them. The cost of making ribosomes is consider-

able. In exponentially-growing bacterial cells, about 80%

of transcriptional activity can be accounted for by pre-

rRNA synthesis (most of the remainder being tRNA syn-

thesis) and up to 25% of protein synthesis activity is dedi-

cated to making r-proteins (Bremer and Dennis , 1987 ). The

situation appears similar in growing yeast, where 60% of

total transcription is of rRNA and, in addition, 50% of the

newly-made mRNAs code for ribosomal proteins (Warner ,

1999 ). Compared to the expenditures for rRNA synthe-

sis, the cost of making mRNAs is negligible. Indeed, the

number of active ribosomes is growth-limiting for a wide

variety of single-celled organisms, bacterial and other

(Scott et  al. , 2010 ; Ehrenberg et  al. , 2013 ). This suggests

that ribosome production could be maximized for speed

and that ribosomal degradation could directly slow cell

growth.

There are, however, three reasons why degrading ribo-

somes might be a good idea at least some of the time. First,

the assembly process is complex with numerous possibili-

ties for introducing errors. Each Escherichia coli ribosome

consists of two large rRNAs, one medium-size rRNA and

about 54 carefully assembled r-proteins; eukaryotic ribo-

somes being more complex still. Considering that the syn-

thesis of individual ribosomal components is unlikely to

be completely stoichiometric and their assembly involves

inefficiencies, timely degradation of ribosomal precursors

stuck on some unproductive sidetrack of their assembly

landscape is likely to be beneficial. Second, because ribo-

somes are normally long-lived, random chemical damage

may accumulate and lead to a gradual loss of function.

These errors can either be rectified by ribosomal repair or

degradation. Third, while a large number of ribosomes are

required for rapid cellular growth, during stress and star-

vation they may be viewed as readily usable repositories

of building blocks for new proteins and nucleic acids; one

cannot grow tomorrow if dead today.

In this review we will sift through current literature rel-

evant for these three scenarios of ribosome degradation,

including references to eukaryotic systems. In addition to

the why of ribosome degradation, molecular mechanisms

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846      Ü . Maiv ä li et al.: Ribosome degradation

of degradation will not be neglected. Emphasis is given to

recent research as there are excellent reviews of the clas-

sics (Deutscher , 2003, 2006, 2009 ).

Degradation of misassembled pre-ribosomes Assembly of ribosomal subunits begins co-transcription-

ally. In E. coli the ribosomal RNAs are synthesized from

5500 nt operons, of which there are seven copies (there

is an extra, eighth, copy of the 5S rRNA gene). As a first

step, this long transcript is cleaved to pre-16S rRNA, and

pre-23S rRNA molecules by the double-stranded RNA-spe-

cific endonuclease ribonuclease III (RNase III) and these

are consequently trimmed to full-length rRNAs by RNase

E, RNase G, RNase T, RNase PH and YbeY (Kaczanowska

and Ryden -Aulin, 2007 ; Deutscher , 2009 ; Davies et  al. ,

2010 ; Shajani et al. , 2011 ; Gutgsell and Jain , 2012 ). Analo-

gously, pre-5S rRNA is created by a digestion at each end

by RNase E. The binding of r-proteins to precursor rRNA is

already required for correct cleavage of pre-23S rRNA by

RNase III (Allas et al. , 2003 ). During the assembly process,

three ribosomal RNAs meet about 54 r-proteins, the rRNAs

undergo at least 38 separate post-transcriptional modifi-

cations and 11 r-proteins accrue one or more modifications

of their own (Kaczanowska and Ryden -Aulin, 2007 ). The

compositions of exponential growth phase and stationary

phase ribosomes are slightly different: stationary phase

ribosomes contain an extra r-protein (S22), a different

isoform of the L31 protein, and increased amounts of the

acetylated L7 as part of the L7/L12 complex (Ramagopal

and Subramanian , 1974 ; Wada , 1998 ; Nanamiya et  al. ,

2004 ). The functional consequences of these differences

remain largely unexplored.

Ribosomal assembly can be mimicked in vitro by

mixing rRNAs with purified r-proteins and incubating the

mixture over extended periods of time under different

non-physiological temperatures (Nierhaus and Dohme ,

1974 ). In living cells the process is relatively fast and

catalyzed by over 20 extra-ribosomal proteins, includ-

ing RNA helicases, ribosome-dependent GTPases, RNA,

and protein chaperones (Kaczanowska and Ryden -Aulin,

2007 ; Shajani et al. , 2011 ). Although the assembly process

is not understood in detail, it is likely that it occurs via

multiple parallel pathways each of which contain rate-

limiting intermediates and unproductive side-paths

(Shajani et al. , 2011 ). A main function of the extra-riboso-

mal assembly factors is to rescue partly assembled parti-

cles from the kinetic traps and cul-de-sac ’ s of the assembly

landscape, thus providing them with new opportunities

to find paths that lead to functional ribosomes. Other

possible functions for assembly factors include blocking

of premature translation by competition for the binding

sites of substrates for translation initiation, blocking

the premature formation of native structural elements

and premature binding of specific r-proteins, as well as

the stabilization and destabilization of pre-ribosomes

(Shajani et al. , 2011 ; Karbstein , 2013 ). In a classic work,

Gausing compared the relative synthesis and accumu-

lation rates of rRNA at different growth rates, which

enabled her to estimate both assembly efficiencies and

rRNA degradation (Gausing , 1977 ). Degradation appears

negligible during rapid growth, while during very slow

growth only about one-third to half of the newly-made

rRNA is incorporated into stable ribosomes. Using similar

methods, upon nutritional down-shift it was shown that

about 50% fewer ribosomes accumulated as mature

particles than were made as rRNAs (Molin et  al. , 1977 ).

These results imply widespread co-assembly ribosomal

precursor degradation in E. coli . Accordingly, RNA puri-

fied from fast-growing E. coli degradosomes consisted

predominantly of rRNA fragments while lacking tRNA

(Bessarab et al. , 1998 ). The degradosome, which consists

of the endonuclease RNase E, the 3 ′ – 5 ′ exonuclease poly-

nucleotide phosphorylase (PNPase), the DEAD-box RNA

helicase RhlB, and the glycolytic enzyme enolase, was

recently shown to interact with mature ribosomes (Tsai

et al. , 2012 ). In addition to PNPase, which is implicated in

ribosome degradation (Basturea et al. , 2011 ), RNase E is

involved in pre-rRNA processing, and is the major RNAse

in mRNA degradation. Elucidation of the functional role

of degradosome-ribosome interaction awaits further

study. It stands to reason that the RNase E endonuclease,

which is readily inhibited by the multitude of structured

elements in correctly-folded rRNA, could nevertheless

recognize and cut misfolded rRNAs that are expected to

have more single-stranded regions. Degradation of mis-

assembled rRNA is likely to be initiated by an endonucle-

ase and to require polyadenylation of degradation inter-

mediates (Maes et al. , 2011 ), which are in turn substrates

for PNPase and RNase R (Basturea et al. , 2011 ). Polyade-

nylation also plays a pivotal role in the degradation of

misassembled tRNA molecules with the participation of

PNPase (Li et al. , 2002 ).

In eukaryotes, rRNAs are synthesized from operons

as in bacteria, but they are longer, are processed in more

complex pathways, are more heavily modified and associ-

ate with about 80 different r-proteins. Accordingly, in yeast

there are around 200 extra-ribosomal assembly factors to

expedite the process (Henras et al. , 2008 ). It seems that as

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Ü . Maiv ä li et al.: Ribosome degradation      847

QC 2

Nob1p

QC 3Exosome

ProteasomeRepair?

RNase T2

Exosome

QC 1

Nucleolus

No-body

16S 23S 5S

Assemblyintermediates

QC 1Exosome, PAP

QC 2YbeY, RNase R

Matureribosomes

YbeY

QC 3

Repair?

100S70S-YbiA70S-YgjD

RNases II, PH, R

Transcription of rrn operon

P body

Ribophagy110S

Recycling

Assembledparticles

Active ribosomes

Recycling

Stress Stress

18S 5,8S 28S

30S 50S 40S60S

60S

40S

ExosomeTRAMP

Ribosomedegradation

Bacteria Yeast

Figure 1   A comparative scheme of Escherichia coli (left) and yeast (right) ribosomal metabolism.

QC 1: The first quality control pathway that degrades ribosomal assembly intermediates using exosome, bacterial poly(A) polymerase (PAP)

and the yeast nuclear poly(A) polymerase complex (TRAMP). QC 2: The second quality control pathway that uses the last processing step of

the 3 ′ -end of the small subunit RNA as a mark for successful assembly of the small subunit. QC 2 requires association of the small subunit

with the large ribosomal subunit to function. QC 3: The third quality control pathway, Non-functional Ribosome Decay (NRD) removes

structurally-sound but functionally-deficient mature ribosomes. In yeast, the inactive 40S and 60S subunits are degraded independently by

different mechanisms. 40S NRD is localized in P bodies and uses the cytoplasmic exosome. The 60S NRD is initiated by r-protein degrada-

tion and occurs in the cytoplasm. The existence of bacterial QC 3 is unclear. Recycling: degradation of ribosomes in response to stress

(including starvation). 100S, 110S, 70S – YbiA and 70S – YgjD denote different inactive ribosomal complexes that are used for ribosome

storage during the stationary growth phase.

the number of components that have to be co-assembled

increases from bacteria to yeast by about 1.5-fold the com-

plexity of the task, as measured by the number of aux-

iliary factors needed for speedy assembly, increases by

about 10-fold. The assembly process starts co-transcrip-

tionally in the nucleolus, where assembly intermediates

can be degraded by the nuclear exosome and continues

after transport at the rate of 2000 ribosomes per minute

to the cytoplasm (Warner , 1999 ; Allmang et  al. , 2000 ).

Pre-ribosomes that are in principle able to participate in

(and potentially to interfere with) protein synthesis are

transported to the cytoplasm in an inactivated form with

several trans-factors bound to active sites, thus preclud-

ing initiation of translation (Panse and Johnson , 2010 ).

After successful transport of pre-ribosomes to the cyto-

plasm there is a final quality control checkpoint of 40S

subunits (Figure 1 ). It seems that 40S subunits that still

contain pre-18S rRNAs lacking the final 3 ′ -end processing

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848      Ü . Maiv ä li et al.: Ribosome degradation

are assembled into translationally inactive 80S ribosomes

solely for quality control purposes (Lebaron et al. , 2012 ;

Strunk et al. , 2012 ). Kinetic competition between the final

processing of pre-18S rRNA by the Nob1p endonuclease

and degradation of the ribosome by unknown means will

ensue (Strunk et al. , 2012 ). In order for the final matura-

tion of the 18S rRNA to occur, the pre-40S must be able

to correctly bind the 60S and participate in the activation

of its GTPase center. The mature 3 ′ -end of the 18S rRNA

would thus serve as a mark that signifies the structural

soundness of the 40S particle. A strikingly similar quality

control mechanism was recently described in E. coli (Jacob

et al. , 2013 ). In this mechanism, the final processing of the

3 ′ -end of pre-16S rRNA is dependent on 70S ribosome for-

mation. Both the ensuing processing step and the alter-

native (ribosome degradation) involve the YbeY protein,

thus making it a prime suspect for the endonuclease that

triggers ribosomal degradation in E. coli . Interestingly,

ribosomes were not only degraded in the absence of

pre-16S rRNA 3 ′ -end processing, but also when ribosomal

structure was perturbed in the presence of the antibiotic

kasugamycin (Jacob et al. , 2013 ). This rhymes with studies

showing that several other ribosome-binding antibiot-

ics can lead to both defective ribosomes and destabiliza-

tion of rRNA (Silvers and Champney , 2005 ; Frazier and

Champney , 2012 ) and suggests a general role for the

YbeY quality control pathway in dealing with structurally

unsound ribosomes.

As in bacteria, misassembled eukaryotic ribosomes

are expected to be scavenged for parts. Interestingly,

all 11 3 ′ – 5 ′ exonucleases of the yeast nuclear exosome

were genetically linked to multiple indirect roles in

various early endonucleolytic pre-rRNA processing steps

(Allmang et al. , 2000 ). As all direct action in these pro-

cessing steps is taken by endonucleases, the most likely

role for the exosome was thought to be in degrading mis-

assembled ribosomal precursor particles (Allmang et al. ,

2000 ). Nuclear pre-ribosomes destined for degradation

are localized to a specific sub-nucleolar compartment

(the No-body), are polyadenylated by the TRAMP complex

and degraded on the spot by the exosome (Dez et  al. ,

2006 ). Similar TRAMP-exosome pathway of rRNA degra-

dation seems to be active in the cytoplasm of human cells

(Slomovic et  al. , 2010 ). Six of the yeast exosome com-

ponents are homologous to E. coli RNase PH and one to

RNase R, both of which, together with the degradosome

component PNPase, are implicated in bacterial ribosome

degradation (Zhou and Deutscher , 1997 ; Deutscher , 2009 ;

Basturea et  al. , 2011 ; Frazier and Champney , 2012 ). The

importance of co-assembly quality control is emphasized

by work indicating that failures thereof are associated

with multiple human diseases (Freed et al. , 2010 ; Narla

and Ebert , 2010 ).

Degradation of inactive ribosomes From the motley crew of ribosomal precursor particles

that pass muster at the quality control steps beautiful

ribosomes are born. Some, however, are still poor at their

primary job of making new proteins. Co-assembly quality

control is likely to recognize larger structural defects,

however it is probable that ribosomes with smaller but

still inactivating changes (mutations or chemical damage)

also occur and duly make trouble when entering poly-

somes. Chemical damage to rRNA and ribosome dysfunc-

tion has been associated with the early stages of human

neurodegenerative disease (Ding et al. , 2005 ; Nunomura

et  al. , 2012 ) and with accelerated ribosomal degrada-

tion in yeast (Mroczek and Kufel , 2008 ). Likely actions

against damaged ribosomes include repair and degrada-

tion. Stable cellular RNAs are readily oxidized by H 2 O

2

treatment in E. coli (Liu et al. , 2012 ) and PNPase has been

implicated in coping with chemically-damaged RNA from

E. coli to human cells (Wu and Li , 2008 ; Wu et al. , 2009 ).

While functional repair of chemically-damaged

mRNA and tRNA by the AlkB demethylase has been

described (Ougland et al. , 2004 ), possible repair of rRNA

has so far escaped scrutiny. There is, however, evidence

for the rejuvenation of E. coli ribosomes by exchanging

r-proteins from elderly ribosomes for newly-made pro-

teins (Pulk et  al. , 2010 ). About 10 different r-proteins

were exchangeable in vivo when measured over 3  h in

stationary-phase cultures while in vitro experiments

suggest that several more could be efficiently exchanged

in principle (Pulk et al. , 2010 ). The physiological impor-

tance of r-protein exchange is as yet unknown.

Degradation of functionally-inactive but fully-assem-

bled ribosomal subunits was first discovered in yeast,

where inactivating point mutations in the decoding center

of the 18S rRNA led to degradation of the 40S ribosomal

subunit, and mutations in the 28S rRNA peptidyl trans-

ferase active site led to degradation of the 60S subunit

(LaRiviere et al. , 2006 ). Quality control mechanisms rec-

ognize the damage at the level of 80S ribosomes. Inter-

estingly, only the mutationally inactivated subunits were

degraded while their wild-type binding partners were

stable, leading to a relative depletion of mutant rRNA

(LaRiviere et al. , 2006 ). Mutations at equivalent positions

of E. coli rRNAs do not lead to similar depletion. This fact

prompted a suggestion that E. coli does not degrade its

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Ü . Maiv ä li et al.: Ribosome degradation      849

inactive but its fully-assembled ribosomes (LaRiviere

et  al. , 2006 ). However, as is common in biology, the

truth may turn out to be more complex. A recent paper

suggests that the YbeY endonuclease requires structur-

ally-defective 30S subunits that are assembled into 70S

ribosomes to initiate ribosomal degradation (Jacob et al. ,

2013 ). This not only identifies YbeY as a potential initi-

ating endonuclease for ribosome degradation but also

raises a strong possibility of a nonfunctional RNA decay

(NRD)-like process in E. coli . Strikingly, both the defective

30S subunit and the associated healthy 50S subunit were

degraded (Jacob et al. , 2013 ).

A different conclusion was reached by Zundel et  al.

(2009) , who showed that subjecting dissociated riboso-

mal subunits, but not 70S ribosomes, to cellular extracts

led to moderate degradation of both 16S rRNA and 23S

rRNA over 45-min incubation at 37 ° C (Zundel et al. , 2009 ).

To clarify the presence or absence of nonfunctional rRNA

decay in E. coli , additional experimental effort is required.

The mechanisms of NRD are being increasingly

exposed in the yeast system and appear to be rather

exciting. The first surprise is that small and large sub-

units are degraded by very different mechanisms. Degra-

dation of the defective and consequently translationally-

stalled small 40S subunits utilizes the same cytoplasmic

No-Go Decay (NGD) pathway as degradation of the mRNA

caught in the stalled ribosome (Cole et  al. , 2009 ). The

stalled 80S ribosome is attacked by the NGD effector pro-

teins Hbs1p, Ski7p, Hbs1p, and Dom34p and the major

cytoplasmic 5 ′ – 3 ′ exonuclease Xrn1p. It is likely that the

Dom34p:Hbs1p complex dissociates the 60S subunit

from the stalled translational complex (Tsuboi et  al. ,

2012 ). Subsequently, 18S rRNA is degraded by the cyto-

plasmic exosome localized in P bodies, along with the

mRNA that is degraded by the same pathway (Cole et al. ,

2009 ; Lafontaine , 2010 ).

The second surprise is that inactive large subunits

(60S) are degraded not only by a different mechanism but

that degradation starts from proteins, not rRNA. Initially

the E3 ubiquitin ligase complex ubiquitylates several pro-

teins (Fujii et  al. , 2009 ). The defective 60S subunits are

then dissociated from the 80S ribosomes with the help of

Cdc48 ubiquitin binding complex and 60S degradation is

initiated by the proteasome (Fujii et al. , 2012 ). Only then

does the 28S rRNA degradation commence, possibly cata-

lyzed by the cytoplasmic exosome near the nuclear enve-

lope (and thus not localized to the P bodies) (Cole et al. ,

2009 ). The 40S subunit that was bound to the mutant 60S

subunit is likely to dissociate from mRNA and to be reuti-

lized in a next round of initiation, therefore avoiding the

need to degrade it via the NGD pathway.

Ribosomal degradation in stress There is a downside to the ubiquity of ribosomes. During

many growth conditions ribosome concentrations limit

the growth rates (Scott et  al. , 2010 ) and, therefore, the

more the merrier. However, cessation of growth can mean

trouble. The once rate-limiting protein synthesis now

turns into a more specialized affair, making most ribo-

somes a surplus product of the cellular economy. As with

any surplus, ribosomes can either be stored in anticipa-

tion of better days or recycled. Indeed, there is convincing

evidence that upon entry to the stationary phase many

ribosomes are stored as inactivated 100S dimers, 70S mon-

omers or dissociated subunits, and this process is rapidly

reversible when growth resumes (Agafonov et  al. , 1999 ;

El -Sharoud, 2004 ). Testing several natural E. coli isolates

in batch culture revealed that over half of the ribosomes

are present as 100S at the onset of the stationary phase

(Wada et  al. , 2000 ). Upon continuance of the stationary

phase, these ribosomal dimers mostly dissociated after a

few hours, only to re-engage for several more days. This

was followed by the final dissolution and degradation of

ribosomes concomitant with loss of viability (Wada et al. ,

2000 ). Two proteins, RMF and HPF, are required for 100S

formation. RMF binds to the 30S subunit and blocks anti-

Shine-Dalgarno:Shine-Dalgarno (anti-SD:SD) interaction

between 16S rRNA and mRNA. HPF blocks the tRNA, IF1

and IF3 binding sites (Polikanov et al. , 2012 ).

A second alternative for ribosome storage is by inac-

tivation of the ribosome in the 70S or at the subunit

level. There seem to be several ways to achieve this. For

example, binding of the YfiA protein, which has a par-

tially overlapping binding site with HPF, leads to inac-

tive 70S formation (Polikanov et  al. , 2012 ). A family of

proteins, exemplified by YqjD, inactivate the station-

ary phase ribosomes and tie them to cell membranes in

E. coli (Yoshida et  al. , 2012 ). A third alternative is that

RsfA (also known as YbeB) protein inactivates stationary-

phase ribosomes by binding to the 50S and dissociating

the subunits ( H ä user et al. 2012 ). Recently, stress-induced

reversible ribosomal 110S dimers were found in rat (but

not in mouse or human) cell lines, hinting that multicel-

lular organisms might use similar strategies of ribosomal

storage upon cessation of growth (Krokowski et al. , 2011 ).

While bacterial ribosome dimers are connected by the

small subunits, the mammalian version appeared to be

connected by the 60S large subunits.

In an instance of ribosome reorganization, during

various stress conditions including stringent response,

oxidative stress and heat shock, 43 nucleotides can

be cleaved from the 3 ′ -end of the mature 16S rRNA by

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850      Ü . Maiv ä li et al.: Ribosome degradation

the endonuclease MazF (Vesper et  al. , 2011 ; Moll and

Engelberg -Kulka, 2012 ). This is likely to happen in fully

assembled ribosomes and results in the accumulation

of specific stress ribosomes that lack anti-SD sequences.

Stress ribosomes reprogram global translation pat-

terns giving preference to leaderless mRNAs lacking SD

sequences (Vesper et  al. , 2011 ). Interestingly, the same

MazF endonuclease also cleaves several mRNAs, con-

verting them into a conveniently leaderless form for the

benefit of the stress ribosomes (Vesper et al. , 2011 ).

So far we have looked at ribosome degradation caused

either by defective processing and/or assembly of the pre-

ribosomes or by the defective structure of the mature ribo-

somes. In fact, most studies on ribosomal degradation

over the past 50  years have focused on the degradation

of functionally-healthy ribosomes. This work has focused

on various cellular stress conditions in E. coli , starting

with Mg 2 + starvation (Aronson and McCarthy , 1961 ) and

including starvation of phosphate, nitrogen, and carbon,

growth in a minimal sea-salt medium, overexpression

of an ectopic protein containing many rare codons, and

increased membrane permeability (Kaplan and Apirion ,

1975 ; Davis et al. , 1986 ; Dong et al. , 1995 ; Kalpaxis et al. ,

1998 ; Deutscher , 2003, 2006, 2009 ). These experiments

all have at least one thing in common: degradation is

mea sured in non-growing stressed cells. In physically-

damaged dying bacterial cells, RNase I can move from its

normal periplasmatic location to the cytoplasm and effi-

ciently degrade ribosomes (Deutscher , 2009 ). In plants,

the RNase I homologue, RNase T2, is located in the endo-

plasmic reticulum and vacuoles, and is thought to be

involved in ribosome recycling under normal conditions

(Hillwig et al. , 2011 ), as are several RNase T2 enzymes of

the ciliate Tetrahymena (Andersen and Collins , 2011 ) and

lysosomal RNase T2 enzymes of fish and humans (Mac-

Intosh , 2011 ). In plants and fungi the RNase T2 enzymes

have been proposed to participate in starvation-induced

phosphate recycling from the ribosomes (MacIntosh ,

2011 ). In E. coli there seems to be an active mechanism

by which the ribosomes inhibit RNase I and thus protect

themselves (Kitahara and Miyazaki , 2011 ). Protection

from degradation is provided by sequestering of RNase I

to helix 41 of the 16S rRNA. This protection of ribosomes

from RNase I increased viability in stationary phase and

greatly increased survival upon artificial depolarization

of the inner cell membrane (Kitahara and Miyazaki , 2011 ).

During starvation in E. coli , the cytoplasmic exonucle-

ases RNase PH, RNase II and RNase R are important for

the removal of rRNA fragments while the endonuclease(s)

that initiate rRNA degradation, apart from RNase I, remain

unknown (Deutscher , 2009 ; Basturea et al. , 2011 ).

While the above implies that ribosomal degradation

can kill, interesting data presented by Basturea et  al.

suggest that in some cases degrading most of your ribo-

somes can be life-saving (Basturea et  al. , 2012 ). They

showed that switching E. coli cultures from phosphate to

arsenate led to degradation of the ribosomes. After the

switch, culture growth stopped for about 80 h after which

a small tolerant cell population resumed growth. The sim-

plest explanation is that such growth must use phosphate

released from degraded ribosomes as the major source of

this essential element. This kind of reasoning is echoed in

yeast, where many cellular recycling pathways, including

ribosome breakdown, were proposed to be required for

stress survival (Davey et al. , 2012 ).

Nevertheless, to degrade functional ribosomes the

cells do not have to be facing death. Even growing E. coli batch cultures can exhibit widespread ribosome degrada-

tion that coincides with a slowing of culture growth before

the onset of the stationary phase (Piir et  al. , 2011 ). This

degradation of ribosomal RNA is genetically independ-

ent of the stringent response, which is the major cellular

pathway for coping with nutrient starvation by inhibi-

ting ribosome synthesis and generally reorganizing gene

expression (Dalebroux and Swanson , 2012 ). Interestingly,

ribosomes were stable during both constant rate growth

and in the actual stationary phase (Piir et  al. , 2011 ). A

similar degradation pattern of ribosomes during prepara-

tion for stationary phase entry was previously described

in various Salmonella strains, where rRNA is normally

fragmented by specific RNase III cleavages and yet consti-

tutes fully functional ribosomes (Hsu et al. , 1994 ). Riboso-

mal degradation that precedes the stationary phase could

either represent an active coping strategy or be an unfor-

tunate cause for the cessation of growth. In this respect,

the emerging picture of the regulation of a major rRNA

degrading enzyme, RNase R, is of note (Figure 2 ). The

E. coli RNase R protein is normally extremely unstable

with a half-life of about 10  min (Chen and Deutscher ,

2010 ). During cold shock, growth on minimal medium

and entry into stationary phase the protein is stabi-

lized so that, even while its mRNA levels are reduced,

its level increases by several-fold (Chen and Deutscher ,

2005 , 2010). In exponentially growing cells, the RNase R

protein is acetylated at a single Lys-residue by the lysyl

acetyltransferase (Pka), which in turn is absent from

late-exponential phase and stationary phase cells (Liang

and Deutscher , 2011 ). The presence of the acetylation is

required for the binding of the tmRNA-SmpB complex to

RNase R (Liang and Deutscher , 2012a ), which in turn leads

to degradation of RNase R by the HslUV and Lon proteases

(Liang and Deutscher , 2012b ). As the well-established

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Ü . Maiv ä li et al.: Ribosome degradation      851

function of the tmRNA-SmpB complex is to rescue stalled

ribosome-mRNA complexes, it stands to reason that under

conditions where more ribosome stalling occurs less

tmRNA-SmpB will be available to destabilize RNase R and

this could lead to increased ribosome degradation. While

RNase R is stabilized in the stationary phase, the Lon pro-

tease is activated by the accumulation of polyphosphate

concomitant with stationary phase onset (Ault -Rich é

et al., 1998 ; Kuroda et al. , 2001 ). The Lon-catalyzed pro-

teolysis upon nutritional downshift leads to degrada-

tion of about 15% of cellular protein, including partial

degradation of most ribosomal proteins (Kuroda et  al. ,

2001 ). This proteolysis activity generates a fresh pool of

reusable amino acids and is necessary for E. coli growth

during nutritional downshifts (Kuroda et al. , 1999, 2001 ).

Polyphosphate synthesis is stimulated by ppGpp, suggest-

ing a role for stringent response in the degradation of ribo-

somal proteins (Kuroda et al. , 1997 ), if not for ribosomal

RNA (Piir et al. , 2011 ).

Similarly, upon ribosome degradation in yeast, most

r-proteins are degraded rapidly while the ones that have

extraribosomal functions or are otherwise required in a

free cytoplasmatic pool may use specific mechanisms of

stabilization (Nusspaumer et al. , 2000 ; Jia et al. , 2012 ).

It is interesting to compare these bacterial processes

with active ribosome degradation in eukaryotes. In yeast

growing rapidly in a nutrient rich medium, cytoplasmic

ribosomes are protected from degradation by the ubiquitin

ligase Rsp5 (Shcherbik and Pestov , 2011 ). When nutrients

become limiting, ribosomal degradation commences in

yeast cultures that continue to grow (Johnston et al. , 1977 ;

Ju and Warner , 1994 ). The magnitude of degradation

appears to be controlled by the ribosome-associated

protein Stm1p to allow for enough remaining translational

capacity for optimal outgrowth upon nutrient upshift

(Van Dyke et al. , 2013 ). During nitrogen starvation, despite

the fact that both RNA and protein synthesis continue, no

net accumulation of RNA due to concomitant RNA deg-

radation occurs (Johnston et al. , 1977 ). Thus a surprising

picture emerges: during nutrient limitation the synthesis

of ribosomes is accompanied by concomitant degrada-

tion. This seems to hold true for both bacteria and yeast.

In yeast, mature ribosomes are degraded on nutrient

limitation by a selective type of autophagy (ribophagy),

which requires the Ubp3/Bre5 ubiquitin protease (Kraft

et  al. , 2008 ; Cebollero et  al. , 2012 ). This pathway is not

inhibited by the Rsp5 ubiquitin ligase (Shcherbik and

Pestov , 2011 ). Ubp3 Δ cells are not only sensitized to nutri-

ent limitation but also to rapamycin, a drug that inhibits

the central target of rapamycin (TOR) signaling pathway

(Kraft et al. , 2008 ). Inhibition of the TOR pathway leads to

growth inhibition through the global inhibition of protein

synthesis, inhibition of the synthesis of new ribosomes,

as well as leading to a large-scale degradation of cytoplas-

mic ribosomes (Pestov and Shcherbik , 2012 ). Although

rapamycin-induced degradation requires the cytoplasmic

exosome for scavenging of rRNA degradation intermedi-

ates, it appears to be genetically distinct from non-func-

tional ribosome decay and from the ribophagy pathway

(Pestov and Shcherbik , 2012 ). It is therefore likely that

this type of ribosome degradation utilizes a yet another

uncharacterized molecular pathway.

Pka

Starvation: slowing growthExponential growth

Ac

E P A 50S

30S

Polyphosphate

AcAc

rRNA fragments

Lon

r-proteinsrRNA deg

RNase Rdeg

r-proteins

SmpB-tmRNA

Rnase R

Figure 2   Regulation of starvation-induced ribosomal degradation in Escherichia coli . During fast growth, RNase R is acetylated by Pka, the acetylated form binds SmpB-tmRNA, which in turn recruits the Lon protease to

degrade RNase R. Starvation represses Pka expression leaving stabilized RNase R free to degrade ribosomes. During starvation, SmpB-

tmRNA binds to stalled ribosomes, terminates the nascent peptide and helps to dissociate the ribosomal subunits. Dissociated subunits

are susceptible to degradation. While ensuing rRNA fragments are scavenged by RNase R and RNase PH, many basic r-proteins are likely to

bind to highly-charged polyphosphate, which is synthesized in the stationary growth phase, and is subsequently degraded by Lon, which

also binds to polyphosphates.

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852      Ü . Maiv ä li et al.: Ribosome degradation

In addition to the control of cellular proliferation,

ribosome degradation is associated with apoptosis from

yeast to humans (King et  al. , 2000 ; Hoat et  al. , 2006 ;

Mroczek and Kufel , 2008 ), diabetes in rats (Ashford and

Pain , 1986 ) and the colony collapse disorder in honey

bees (Johnson et al. , 2009 ).

Future directions The relative plentitude of situations where ribosomes are

degraded in conjunction with the number of different

degradation pathways that seem to be required lead to the

happy conclusion that there are things yet to look forward

to in the ribosome degradation field. We still know very

little about the mechanisms that trigger the various ribo-

somal quality control pathways. A glaring omission, that

has just begun to be looked into, is the identities and

modes of action of the endonucleases that initiate ribo-

somal degradation. The presence and possible nature of

a NRD-like pathway in bacteria clearly warrants further

study. We believe that as ribosomal degradation is but

one solution to various problems arising in the cellular

economy, this process cannot be fully understood without

sufficient understanding of the catalysis of ribosome

assembly, repair and storage.

Now that instances of degradation of the most ubiqui-

tous stable RNAs, including tRNAs, are revealed in increas-

ing numbers (Thompson and Parker , 2009 ; Wilusz et  al. ,

2011 ; Dewe et  al. , 2012 ), a rather obvious question arises:

how stable are the other stable ribonucleoproteins ? These

include the RNase P ribonucleoprotein, the various splice-

osomes, and small nucleolar ribonucleoproteins involved

in RNA modification, and others, all of which could benefit

from quality control and regulation at the level of degrada-

tion. Lessons learned in the ribosome could thus help to shed

new light to superficially-unrelated biological questions.

Acknowledgements: We thank Aivar Liiv and David

Schryer for helpful comments on the manuscript and Sille

Hausenberg for help with preparing the figures. This work

was supported by the Estonian Science Agency grant no

9040 and by the European Regional Development Fund

through the Center of Excellence in Chemical Biology.

Received February 7, 2013; accepted March 20, 2013; previously

published online March 24, 2013

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Ülo Maiväli is a molecular biologist studying ribosomal metabolism

in Escherichia coli . He obtained his PhD from the University of Tartu

in 2004. Currently he is a researcher in the Institute of Technology,

University of Tartu, Estonia.

Anton Paier has a Master’s degree in Modern Literature from the

University of Genua, Italy, a Bachelor’s in Military History from the

University of Stockholm and a Master’s in Biomedical Laboratory

Sciences from the Karolinska Institute (2008). Currently he is doing

a PhD in Molecular Biology at the University of Tartu. His thesis is

centered on the ribosomal degradation in E. coli .

Tanel Tenson is a biochemist and microbiologist studying the

mechanisms of antibiotic action and antibiotic resistance. He

obtained his PhD from the University of Tartu in 1997. Tanel Tenson

is currently Professor of Technology of Antimicrobial Compounds at

the Institute of Technology, University of Tartu.

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