molecular mechanisms of the prl phosphatases

REVIEW ARTICLE

Molecular mechanisms of the PRL phosphatasesPablo Rios, Xun Li and Maja Kohn

European Molecular Biology Laboratory, Genome Biology Unit, Heidelberg, Germany

Keywords

cancer; cell signalling; dual specificity

phosphatases; metastasis; phosphatase of

regenerating liver; protein phosphatases

Correspondence

M. Kohn, European Molecular Biology

Laboratory, Genome Biology Unit,

Meyerhofstrasse 1, 69117 Heidelberg,

Germany

Fax: +49 6221 387518

Tel: +49 6221 3878544

E-mail: [email protected]

(Received 8 January 2012, revised 20

February 2012, accepted 9 March 2012)

doi:10.1111/j.1742-4658.2012.08565.x

The phosphatases of regenerating liver (PRLs) are an intriguing family of

dual specificity phosphatases due to their oncogenicity. The three members

are small, single domain enzymes. We provide an overview of the phospha-

tases of regenerating liver, compare them to related phosphatases, and

review recent reports about each phosphatase. Finally, we discuss similari-

ties and differences between the phosphatases of regenerating liver, focus-

ing on their molecular mechanisms and signalling pathways.

Common features of the PRLs

Ever since phosphatase of regenerating liver (PRL)-3

was found to be overexpressed in liver metastatic tissue

originating from colon cancer, but not in normal colon

tissue nor in the primary tumour [1], the PRL family

of phosphatases has received much attention. There is

strong evidence suggesting that not only PRL-3, but

also PRL-1 and PRL-2 are oncogenes and, as such,

belong to the few phosphatases that lead to the devel-

opment of cancer [2–5]. The PRLs promote cell prolif-

eration, migration, invasion, tumour growth and

metastasis [4,6–15], and these are recognized driving

forces behind their oncogenicity. The underlying

molecular mechanisms still remain undetermined,

although progress has been made in understanding the

proteins and pathways involved [2,3].

Rat PRL-1 phosphatase was the first member of the

PRL family to be discovered, and was found as an

immediate early gene induced in rat regenerating liver

and mitogen-stimulated cells, and to be constitutively

expressed in insulin-treated rat hepatoma H35 cells

[16,17]. Subsequently, human PRL-2 was identified in

a genetic study [18] and, together with human PRL-1,

in a prenylation screen as farnesylated proteins in vitro

through a C-terminal CAAX motif (where C is the

Abbreviations

AML, acute myeloid leukaemia; ATF, activated transcription factor; bGGT II, b-subunit of Rab geranylgeranyltransferase II; CDC14, cell

division cycle 14 homologue; CDK2, cyclin-dependent kinase 2; Csk, C-terminal Src kinase; EF-2, elongation factor 2; EMT, epithelial–

mesenchymal transition; ERK, extracellular signal-regulated kinase; FAK, focal adhesion kinase; FKBP38, FK506-binding protein 38; GTPase,

guanosine triphosphate phosphohydrolase; KAP, kinase-associated phosphatase; MDM2, mouse double minute 2; MEF, mouse embryonic

fibroblast; MEF2C, myocyte enhancer factor 2C; MEKK1, mitogen-activated protein ⁄ extracellular signal-regulated kinase kinase kinase 1;

MMP, matrix metalloproteinase; OMFP, ortho-methylfluorescein phosphate; PCBP1, polyC-RNA-binding protein 1; PI3K, phosphatidylinositol

3-kinase; PI(4,5)P2, phosphatidylinositol 4,5-bisphosphate; PIRH2, protein with a RING-H2 domain; pNPP, para-nitrophenyl phosphate;

PRL, phosphatase of regenerating liver; PTEN, phosphatase and tensin homologue; PTP, protein tyrosine phosphatase; PTPMT1, protein

tyrosine phosphatase mitochondrial 1; rhoGAP, Rho-GTPase-activating protein; SH3, SRC homology 3 domain; siRNA, small interfering RNA;

Src, sarcoma; TGF, transforming growth factor; VEGF, vascular endothelial growth factor; VHR, vaccinia H1-related phosphatase.

FEBS Journal (2012) ª 2012 The Authors Journal compilation ª 2012 FEBS 1

cysteine that is prenylated, A is an aliphatic amino

acid and X is any amino acid) [6]. In addition, the lat-

ter study already recognized the oncogenic potential of

these phosphatases. Finally, mouse PRL-2 and PRL-3

were cloned and analyzed with respect to their

sequence similarity to other phosphatases and the

expression pattern in mice [19]. Although the amino

acid identities of the PRLs are low compared to other

phosphatases, they are very high between the three

PRLs: 87% between PRL-1 and PRL-2; 79% between

PRL-1 and PRL-3; and 76% between PRL-2 and

PRL-3 in humans [2,3,20] (Fig. 1).

PRLs are classified into the family of dual specificity

phosphatases (DSP) (also called vaccinia H1-like phos-

phatases) [21], which is a subgroup of the class I pro-

tein tyrosine phosphatase family defined by the

conserved active site p-loop sequence HC(X)5R[S ⁄T][22]. PRLs are relatively small proteins of approxi-

mately 20 kDa. They do not have regulatory domains,

although they contain a variety of intrinsic regulatory

elements [2,3]. PRLs are the only phosphatases of the

protein tyrosine phosphatase (PTP) superfamily that

carry the aforementioned CAAX motif and are

farnesylated in vivo [23–25], and they can also be gera-

nylgeranylated in vitro [6,23,26]. However, reports

about PRL-3 geranylgeranylation are contradictory

[23,26]. Interestingly, the CAAX box is common

amongst human phosphatidylinositol-5-phosphatases

[27]. Prenylation of the PRL phosphatases localizes

them to the plasma membrane and intracellular

membranes in distinct punctate structures, which are

suggested to be the early endosome, and deletion of

the CAAX box or application of farnesyl transferase

inhibitors prevents membrane localization and

redirects the PRLs into the nucleus [10,23,28]. The cat-

alytic activity and farnesylation are necessary for the

cellular and tumour- and metastasis-related phenotypes

of the PRLs [8,28–30]. In addition, similar to members

of the Ras superfamily of guanosine triphosphate

phosphohydrolases (GTPases), the PRLs carry a

Fig. 1. Structure-based multiple sequence alignment of PRLs and the structurally most closely-related PTPs. Ci-VSP is included as a result

of its activity toward PI(4,5)P2 [48] and PTPMT1 as a result of its activity toward PI(5)P [47]. The PRLs are depicted in full length; other PTPs

are shown in truncated versions according to relevance with respect to sequence alignment with the PRLs. The amino acids are colored by

polarity: A, P, V, I, W, F, L, G, M (black); S, T, Y, N, C, Q (green); D, E (red); H, K, R (blue). The consensus residue and conservation rate at

each position are shown below the sequences. An ambiguous residue is indicated as ‘X’. The putative ‘CXnE’ motif, the WPD-loop and the

active site p-loop are indicated in red squares. Protein sequences were manually associated with 3D homologous structures in STRAP

(http://3d-alignment.eu/) and the alignment was computed with CLUSTALW_3D. The alignment was manually adjusted according to the

superimposed structures.

Molecular mechanisms of the PRL phosphatases P. Rios et al.

2 FEBS Journal (2012) ª 2012 The Authors Journal compilation ª 2012 FEBS

polybasic region adjacent to the CAAX motif, which

aids in mediating the membrane localization of PRLs

[10,31,32] and could be involved in mediating nuclear

localization [10,25], although probably not as a

bipartite nuclear localization sequence [33].

As shown in vitro and in cells for PRL-1, as well as

in vitro for PRL-3, another regulatory feature could be

the formation of trimers or other oligomers

[10,31,33,34]. Indeed, PRL-1 crystallized in trimers

[31,34]; however, the NMR structures of PRL-3

revealed a monomeric state [32,35]. This discrepancy

could be a result of the different experimental meth-

ods, although it could also mean that there are differ-

ences within the PRL family with respect to the

ability ⁄ tendency to form oligomers. PRL-1 oligomeri-

zation, which requires C-terminal farnesylation, was

reported to be necessary for its function in cells [10].

By contrast, for PRL-3, it was shown that farnesyla-

tion-dependent oligomerization decreased the in vitro

phosphatase activity toward an unnatural substrate

[33]. Owing to the more complex environment in cells,

it is likely that the measured in vitro activity of PRL-3

does not reflect the in vivo behaviour.

The structural elucidation of PRL-1 and PRL-3 also

revealed the importance of cysteine 49. This cysteine is

localized very closely to the active site cysteine in all

structures and the two cysteines can form a disulfide

bond [31,32,34,36]. As for other PTPs, this indicates

that PRLs are subject to redox-regulation, following

the same mechanism as other DSPs such as phospha-

tase and tensin homologue (PTEN) [37]. It was specu-

lated that this redox-mechanism not only regulates the

activity of the phosphatases, but also could protect the

catalytic Cys104 from further and irreversible oxida-

tion [31,32]. Skinner et al. [38] reported that the reduc-

tion potential of this disulfide bond for PRL-1 in vitro

is lower ()365 mV) than the reduction potential range

in normal cellular environments ()170 to )320 mV),

indicating that newly-synthesized PRL-1 in cells could

be oxidized and thereby inactive. Interestingly, the

same study reported that the C-terminal CAAX box

farnesylation motif (CCIQ in PRL-1) also regulates

the PRL-1 activity in vitro. When mutating the C170

and C171 residues of the CAAX motif, the resistance

to oxidation of PRL-1 was increased, mediated by con-

formational changes. Such a conformational switch

would likely also occur upon farnesylation, increasing

the catalytic activity as a result of a lower sensitivity

to oxidation. Thus, farnesylation could not only regu-

late PRL-1 subcellular localization, but also the cellu-

lar functions that are dependent on catalytic activity

[38]. Pascaru et al. [33] reported that CAAX deleted

PRL-3 also displays enhanced catalytic activity

compared to wild-type PRL-3, which suggests that the

C-terminus and farneslyation are common features for

regulating the catalytic activity of the PRL proteins.

As similar as the PRLs may appear, many intriguing

differences are already apparent. In the present review,

we first compare the structural features of the PRLs to

related phosphatases and then describe the features for

each PRL in order of how well studied they are, focus-

ing on a discussion of key characteristics and signalling

pathways, as well as the novel insights that have

appeared subsequent to previous reviews [2,3]. Next,

we discuss the differences between the PRLs. Finally, a

discussion about the catalytic site architecture of the

PRLs and its putative influence on the substrate recog-

nition mechanism is provided.

Comparison of structural featuresof the PRLs with other relatedphosphatases

In general, structural analyses revealed that the PRLs

have hydrophobic, shallow binding pockets and a wide

pocket entrance [31,32,34,35]. Compared to the struc-

turally most closely-related phosphatases [PTEN,

vaccinia H1-related phosphatase (VHR), cell division

cycle 14 homologue (CDC14), kinase-associated phos-

phatase (KAP)], PRLs lack helices and loops that can

be important for substrate recognition [31,34]. In addi-

tion, all of those phosphatases, which are structurally

most closely related, have very different substrate spec-

ificities, ranging from pTyr (VHR) [39], pSer (CDC14)

[40] and pThr (KAP) [41] to phosphatidyl inositol

phosphates (PTEN) [42], making direct conclusions

with respect to the substrate specificity of PRLs from

this general comparison impossible. For a more

detailed comparison, in Fig. 2 the crystal structure of

the catalytic pocket of PRL-1 complexed with a sulfate

ion [34] is overlaid with complexed structures of the

related phosphatases PTEN [43], VHR [44], CDC14

[45] and KAP [46], and also with PTP mitochondrial 1

(PTPMT1) [47] and the PTEN-like phosphatase

Ci-VSP from Ciona intestinalis [48,49]. The latter two

were chosen due to their ability to dephosphorylate the

5-position of phosphatidylinositol phosphates in light

of the fact that PRL-3 has been proposed to be a

phosphoinositide-5-phosphatase (see below) [50]

(unfortunately, complexed structures of PRL-3 are not

yet available). In general, the active site of PRL-1

overlays well with all of the structures. The best

matches appear to be KAP and CDC14B, whereas the

worst overlay is with VHR due to the many different

amino acids. Thus, if a conclusion can be drawn from

this comparison, PRL-1 likely prefers pThr ⁄pSer

P. Rios et al. Molecular mechanisms of the PRL phosphatases


residues similar to KAP and CDC14B. Nevertheless, it

was proposed that significant structural rearrange-

ments will likely occur upon association of PRL phos-

phatases with their physiological substrates [31], and

thus only structures of PRLs with their substrates are

able to answer the question of how these interactions

look like.

Intriguingly, PRLs do not contain the conserved

[Ser ⁄Thr] residue of the PTP active site p-loop but,

instead, an alanine is found in that position (Fig. 1). It

is thought that this alanine results in the low intrinsic

in vitro activity of the enzymes [31,32] due to the role

of this Ser ⁄Thr in the catalytic mechanism of PTPs,

aiding in the release of the phosphate from the phos-

phatase [51]. Replacement of alanine with serine

enhanced the catalytic activity towards unnatural sub-

strates such as para-nitrophenyl phosphate (pNPP) or

ortho-methylfluorescein phosphate (OMFP) [31,32,50].

This is in agreement with observations for the mouse

phosphatase LDP-2, which also carries an alanine in

the respective position [52]. However, the human

orthologue, DUSP19 (SKRP1) also displayed a low

activity toward pNPP compared to other DSPs, but

this activity was not enhanced when the alanine was

Fig. 2. Structural comparison of active site of PRL-1 bound to a sulfate ion (1XM2, white) with complexed structures of closely-related

PTPs: PTEN (1D5R, green), KAP (1FPZ, cyan), CDC14B (1OHE, yellow) and VHR (1VHR, pink). Ci-VSP (3AWF, blue) is depicted as a result

of its activity toward PI(4,5)P2 [48], and PTPMT1 (3RGQ, orange) as a result of its activity toward PI(5)P [47]. Amino acids are numbered

according to protein database files.



replaced with a serine [53]. Curiously, an Ala to Ser

mutation in PRL-3 completely abolished phosphatase

activity toward a potential natural substrate [50]. Thus,

the role of the natural serine to alanine mutation is

not clear; it was suggested that it could be involved in

substrate recognition [52] or structural integrity [50],

although its role could also be different for every phos-

phatase that carries this mutation.

Another conserved loop in the PTP superfamily is

the WPD loop, of which the Asp acts as general acid

in the catalytic reaction [51]. In the PRL family, this

loop consists of the sequence 68 ⁄ 65WPFDD72 ⁄ 69 (where

the numbering refers to PRL-1 and PRL-3 ⁄PRL-2)

(Fig. 1) and, for PRL-1 and PRL-3, it was shown that

the catalytically active Asp is the D72 [25,32]. It is not

known whether the additional residues in this loop

have a function. However, as there are quite a few

exceptions in the DSP family with respect to the con-

servation of this loop; the conservation might actually

occur mostly in classical PTPs. For example, neither

PTEN, nor CDC14 carry a tryptophan close to the

catalytically active Asp (Fig. 1) and, in the myotubula-

rin family, the W-D motif is localized within the

p-loop [54]. Indeed, a D92A mutation in the WPD

loop in PTEN [55] and D72A in PRL-3 [50] caused

only a partial loss of activity and, for PTEN, it was

shown that the Asp does not act as the general acid in

the first step of the catalysis [55]. Interestingly, PTEN

shows a high sequence similarity in this region to the

PRLs in that the WPFDD of the PRLs aligns with

YPFED in PTEN (Fig. 1).

As noted above, PRLs carry a regulatory cysteine.

Interestingly, a natural mutation of the regulatory cys-

teine 71 in PTEN, which is mutated to tyrosine in

Cowden disease, led to a loss of phosphatase activity

toward the substrate inositol(1,3,4,5)tetraphosphate

[56]. A C49S mutation in PRL-1 led to slightly lower

activity toward pNPP [31]; by contrast, a C49A muta-

tion in PRL-3 did not lead to a change in activity

against the unnatural substrate OMFP [32]. Consider-

ing that an alanine mutation (C49A in PRL-3) intro-

duces a much less drastic change in electrostatic and

steric properties than a tyrosine mutation (C71Y in

PTEN) does, and that activities toward unnatural sub-

strates can sometimes be misleading [50], it is tempting

to speculate whether these regulatory cysteines fulfill

other tasks in the respective phosphatases, such as

maintaining structural integrity or aiding in substrate

recognition, for example through the correct position-

ing of amino acids that are involved in substrate inter-

actions or the catalytic mechanism. This idea is fueled

by another interesting consideration: PTPMT1 con-

tains the catalytically relevant ‘EEYE’ loop, in which

Glu73 and Glu76 were shown to be essential for cata-

lytic acticity and Glu76 interacts with and stabilizes

the conserved catalytic Arg in the catalytic p-loop in

the crystal structure [47]. As shown in Fig. 1, all of the

depicted regulatory cysteine-carrying DSPs have a Glu

close to this Cys, which aligns well with Glu73 and ⁄orGlu76 of the ‘EEYE’ motif either in the backbone

and ⁄or in the side chain (Fig. 3). Even in KAP, where

there is a small loop between the Cys and the Glu, the

alignment is excellent, and also Ci-VSP from a sea

squirt contains a Glu next to a Cys and aligns very

well (Fig. 3). Structurally closely-related DSPs that do

not contain a regulatory cysteine, with the exception

of PTPMT1, do not have a Glu in that position

(Fig. 1). Since mutation of the general acid of the

‘WPD loop’ in PTEN [55] and PRL-3 [50] reduced,

but did not abolish, catalytic activity toward the

(potential) natural substrate, and C71Y mutation

diminished the catalytic activity of PTEN [56], it is

worth investigating whether the acidic amino acid

adjacent to the regulatory cysteine could play an

important role in the general catalytic mechanism or

stabilize the catalytic pocket of DSPs that carry a

putative ‘CXnE’ motif (where X = any amino acid

and n = the number of amino acids between the C

and E residues; e.g. 0 for PRL-1 and PRL-3; 1 for

PTEN and CiVSP; 3 for KAP).

Fig. 3. Structural alignment of the putative ‘CXnE’ motif. The Glu

residues adjacent to the regulatory cysteines in the crystal struc-

tures of PRL-1 (green: 1XM2), PTEN (purple: 1D5R), KAP (pink,

1FPZ) and Ci-VSP (yellow: 3AWF) align well with the Glu of the

‘EEYE’ loop in PTPMT1 (cyan: 3RGQ). The structure of PTEN is in

complex with an inhibitor [L(+)-tartrate], which may be the reason

why the side chain of PTEN does not align with Glu144 of

PTPMT1, although the backbone aligns with the Glu141 of

PTPMT1. Only complexed sructures are compared here. Amino

acids are numbered according to protein database files.



PRL-3

PRL-3 expression, interacting proteins and

regulation

PRL-3 mRNA was found predominantly in the skele-

tal muscle and at moderate levels in the heart, as

shown in mouse [19] and human [57] tissues, and, in

both studies, PRL-3 was also detected in other organs

at lower levels. Interestingly, it was reported that the

expression in the heart only occurs during develop-

ment, and not in the human adult organism, as dem-

onstrated at the mRNA and protein levels [13]. This

finding could have important implications for any

potential drug discovery against PRL-3 because inhibi-

tion of PRL-3 in the adult heart could lead to cardio-

toxic effects [2,20]. Furthermore, PRL-3 was found to

be expressed in the developing blood vessels and pre-

erythrocytes [13], suggesting that PRL-3 plays an

important role in embryogenesis. In addition, Zeng

et al. [23] observed that PRL-3 is present in differenti-

ated villus epithelial cells of the small intestine in mice.

The upregulation of PRL-3 in cancer has received

the most study with respect to the three PRLs, and

was identified in colon [1,58], breast [59], gastric [60]

and ovarian [61] carcinomas. In addition, high levels

of PRL-3 appear to be associated with a poor progno-

ses and, particularly for colon cancer, high levels of

PRL-3 were shown to be predictive for the develop-

ment of liver metastatis [62]. These findings are

reviewed in detail in Bessette et al. [3]. In addition,

PRL-3 was reported to be elevated in oral and cervix

squamous cell carcinomas [63,64]. Furthermore, it was

found to be overexpressed in haematological malignan-

cies, namely in a subset of multiple myelomas [65,66]

and in acute myeloid leukaemia (AML) [67].

A few substrates have been suggested for PRL-3,

namely ezrin [68,69], elongation factor 2 (EF-2) [69],

keratin 8 [70] and integrin b1 [71], all four of which

have been reviewed [2], as well as stathmin [72] and

nucleolin [15]. Recently, we described phosphatidylino-

sitol(4,5)bisphosphate [PI(4,5)P2] as a potential natural

substrate. Although no in vivo activity against PI(4,5)P2

has yet been demonstrated, a correlation between differ-

ences of in vitro activity and phenotype in the cell

migration of wild-type PRL-3 and three PRL-3 mutants

was demonstrated. This correlation was only true

for activity against PI(4,5)P2 and not against the

unnatural substrate ortho-methylfluorescein phos-

phate [50]. Of the putative substrates, direct dephos-

phorylation was demonstrated in the case of ezrin and

PI(4,5)P2, whereas, for EF-2, keratin 8, nucleolin and

stathmin, PRL-3-dependent downregulation of the

phosphorylation level was shown in vivo, and integrin

b1 is now considered to be indirectly affected by PRL-3

[2,73]. In independent experiments, however, the influ-

ence of PRL-3 overexpression on ezrin phosphorylation

could not be confirmed, which may be a result of the

use of different cell lines [70,74]. Stathmin, nucleolin

and keratin 8 were shown to co-immunoprecipitate

with ectopic (inactive) PRL-3, but no direct interaction

with EF-2 was reported. Other direct interaction part-

ners have been identified: integrin a1 [71], cadherin

CDH22 [75] and the peptidyl prolyl cis ⁄ trans isomerase

FK506-binding protein 38 (FKBP38) [76], all discovered

in yeast two-hybrid screens, and PRL-3 itself through

potential oligomerization [10,33]. Most of the proposed

directly interacting proteins are related to the role of

PRL-3 in cell migration and invasion and are connected

in some way to the plasma membrane [ezrin, PI(4,5)P2,

integrin a1, CDH22] or to the cytoskeleton [keratin 8,

stathmin]. Noteworthy, nucleolin is localized to the

cytoplasm and nucleus and is involved in cell prolifera-

tion [15] and FKBP38 is a cytosolic protein that regu-

lates PRL-3 protein levels and proteasomal degradation

in MCF-7 and HCT116 cell lines [76].

In addition, an unbiased mass spectrometry-based

approach revealed 110 potential interacting proteins

when PRL-3 was used as a bait, 38 of which were con-

sidered to be of high confidence [77]. The identified

proteins have not yet been followed up by experimental

validation. It is striking that none of the proposed

binding partners from other studies were identified,

showing how difficult it is to validate substrates and

interacting proteins of PRL-3 (and phosphatases in

general).

PI(4,5)P2 as a substrate for PRL-3 offers a connec-

tion to another substrate, ezrin. Ezrin forms part of

the ERM (ezrin–radixin–moesin) complex, which con-

nects the plasma membrane with the actin cytoskeleton

and is implicated in tumour metastasis [78]. Ezrin

requires PI(4,5)P2 binding and Thr567 phosphorylation

to become active at the plasma membrane [79], so that

it can exert its multiple functions in cell adhesion,

motility, morphogenesis and signalling pathways

[79,80]. In addition to potential PI(4,5)P2 depletion by

PRL-3, PRL-3 is assumed to dephosphorylate Thr567

[68], meaning that PRL-3 could inactivate ezrin in

multiple ways. Considering that the binding of ezrin

by PI(4,5)P2 is required for the phosphorylation of

Thr567, the lower phosphorylation level of Thr567

could also be an indirect effect as a result of the pre-

vention of ezrin binding to the plasma membrane [79].

On the other hand, PRL-3 has been reported to upre-

gulate Src kinase activity [81] (see also below), and Src

can phosphorylate Tyr477 in ezrin, which is required



for anchorage-independent growth and cell invasion in

a 3D environment [82]. Tyr477 phosphorylation was

crucial for the correct localization of ezrin to sub-

membraneous patches in the 3D culture. The influence

of Thr567 phosphorylation and PI(4,5)P2 binding was

not studied in this context; however, other factors

aside from the latter two are important for proper

activity and membrane localization of Ezrin, depend-

ing on the functional context. This shows that the

activity of PRL-3 takes place in a very complex envi-

ronment, which in itself remains incompletely under-

stood.

PRL-3 is subject to complex regulatory mechanisms.

It is known that PRL-3 mRNA levels do not necessar-

ily correspond to protein levels [83] and that PRL-3

abundance is controlled at the transcriptional and

translational levels, as well as through degradation

mechanisms [76] (see above).

PRL-3 is a direct transcriptional p53 target gene in

mouse (mouse embryonic fibroblast; MEF) and human

(H1299 human lung adenocarcinoma, SK-Hep-1 hepa-

tocellular carcinoma) cells [30,84], and ectopic expres-

sion of p53 and p73 increases PRL-3 transcription in

H1299 nonsmall cell lung cancer cells [85]. In other

cancer cells, such as SNU-475, Hep3B and HeLa cells,

the transcriptional level of PRL-3 did not increase

upon ectopic p53 expression [30], suggesting that this

interaction is cell type specific (although two PRL-3

introns harbour a p53 consensus sequence that can

bind the p53 protein) [84]. PRL-3 transcription is also

activated by the vascular endothelial growth factor

(VEGF) through the transcription factor myocyte

enhancer factor 2C (MEF2C) in human umbilical vein

endothelial cells (HUVEC) [86]. MEF2C binds the

promoter region of PRL-3 in vitro and in vivo, and

notably, the presence of MEF2C is critical in heart

and skeletal muscle where PRL-3 is abundant. This,

together with the distinct expression pattern in human

healthy tissues, suggests that transcription of PRL-3

could be controlled by tissue specific transcription fac-

tors [86]. However, an equal enhancement of PRL-3

protein amounts in the presence of MEF2C was not

observed. Interestingly, in PRL-3-positive nonsmall cell

lung cancer cells (NSCLC), elevated levels of VEGF

and its isoform VEGF-C were found, and high levels

of both were correlated with micro and lymphatic ves-

sel density [12], demonstrating that the expression of

PRL-3 facilitates angiogenesis [2].

Snail is a transcription factor involved in the epithe-

lial–mesenchymal transition (EMT). EMT is an impor-

tant process during development and metastasis and,

in this process, cells lose cell–cell adhesion and gain

motility. Snail is known to repress the expression of

E-cadherin, resulting in the disassembly of cell–cell

adhesion junctions and an increase of invasiveness [87].

The overexpression of PRL-3 was demonstrated to

promote EMT, and it was suggested that the action of

PRL-3 leads indirectly to the deinhibition of Snail

[75,88]. Recently, however, Zheng et al. [89] reported

that the PRL-3-encoding gene contains three potential

binding sites of Snail in the promoter region, and that

the transcriptional activity of the PRL-3 promoter was

abolished after the mutation of one Snail binding site.

Snail was suggested to regulate promoter activity and

protein expression of PRL-3 in colorectal cancer cell

lines, which appears to be contradictory to the earlier

reports. Thus, the interaction between PRL-3 and

Snail requires further investigation.

Recently, Jiang et al. [74] reported that PRL-3 is a

direct regulatory target of transforming growth factor

(TGF)b signalling in colon cancer metastasis. TGFbsignalling suppresses the metastasis of colon cancer

cells potentially by inducing stress-induced apoptosis.

It was demonstrated that TGFb signalling inhibited

the expression of PRL-3 in a mothers against decapen-

taplegic homologue (Smad) 3-dependent manner.

Because a loss of TGFb signalling occurs in 30–50%

of colon cancers, this could be a feasible mechanism

for explaining PRL-3 upregulation in colon cancer

[74].

A translational regulator of PRL-3 is polyC-RNA-

binding protein 1 (PCBP1) [83]. PCBP1 overexpression

inhibited PRL-3 expression via interaction with a

GC-rich motif at the 5¢ UTR of PRL-3 mRNA. In

clinical samples of normal and cancerous epithelia, an

inverse correlation between protein levels of PRL-3 and

PCBP1 was observed, and knockdown of endogenous

PCBP1 in HCT-116 cells inhibited tumourigenesis in

mice, indicating that PCBP1 acts as a tumour suppres-

sor in vivo [83].

Signalling pathways affected by PRL-3

The signalling pathways affected by PRL-3 have been

reviewed by Bessette et al. [3] and Al-Aidaroos and

Zeng [2]. Therefore, we only briefly describe the key

signalling effects of PRL-3 and add data that have

appeared subsequent to these reviews.

By demonstrating that PRL-3 upregulates mesenchy-

mal markers and downregulates epithelial markers, it

was shown that PRL-3 promotes EMT [75,88]. It pro-

motes EMT and cell survival by acting upstream of

phosphatidylinositol 3-kinase (PI3K) [74,89]. PI3K

signalling promotes many processes, such as cell

survival, cell proliferation or cell motility, and PI3K is

an oncogene [90,91]. PRL-3 was reported to



post-transcriptionally downregulate PTEN protein lev-

els [88]. PTEN counteracts PI3K activity by converting

phosphatidylinositol triphosphate PI(3,4,5)P3 into

PI(4,5)P2; thus, its downregulation leads to the activa-

tion of PI3K signalling. In addition, PRL-3-mediated

activation of PI3K could relieve the inhibition of the

mesenchymal marker Snail (see above) by inhibition of

glycogen synthase kinase (GSK)-3b [75,88]. Further-

more, PRL-3 was reported to promote cell survival

under growth factor deprivation stress by activating and

maintaining the activity of the PI3K ⁄Akt pathway [74].

PRL-3 was suggested to either reduce the number of

focal adhesions and ⁄or increase focal adhesion turn-

over to mediate cell invasion and motility [2]. Focal

adhesion complexes are multi-component sites where

integrins mediate the contact between the cell and the

extracellular matrix [92]. Levels of PI(4,5)P2 at the cell

membrane are crucial for regulating the dynamics of

focal adhesion complexes [92], and focal adhesion

kinase (FAK) is a key component of focal adhesion

complexes [93]. FAK integrates external signals to pro-

mote cell motility via many different pathways involv-

ing the regulation of (or interaction with) proteins

such as cadherins, Src, p130Cas, Rho-family GTPases

and ezrin [93], many of which were shown to be

affected by PRL-3. Integrin a1 and cadherin-22 were

reported to be direct interactors of PRL-3 (see above),

E-cadherin was shown to be downregulated by PRL-3

[75,88] and PRL-3 signalled via integrin b1 in LoVo

colon cancer cells leading to extracellular signal-regu-

lated kinase (ERK)1 ⁄ 2 activation [73]. Src kinase was

activated by PRL-3 via translational downregulation

of C-terminal Src kinase (Csk), which is a negative reg-

ulator of Src [81,94]. Src activation by PRL-3 led to

the phosphorylation of downstream proteins such as

signal transducer and activator of transcription

(STAT) 3 and p130CAS, and, in agreement with Peng

et al. [81], ERK1 ⁄2. In further studies, PRL-3 acti-

vated RhoC, downregulated Rac-GTP [28,88] and had

no effect on Cdc42 [28]. RhoA activity was reduced by

PRL-3 overexpression in the earlier study [28] and

enhanced in the later study [88]. These findings demon-

strate that the Rho family of GTPases act downstream

of PRL-3, and also show that the regulation is com-

plex. An interesting context in this regard is that active

ezrin recruits both positive and negative regulators of

the Rho family of GTPases [95]. Upon inactivation of

ezrin by PRL-3, these regulators could be released,

which would contribute to maintaining the active form

of the Rho GTPases and may explain the activation of

RhoA and RhoC when overexpressing PRL-3 [2].

The activity of PRL-3 against PI(4,5)P2 [50] offers

the intriguing possibility of PRL-3 regulating all of the

noted proteins upstream of FAK. This regulation,

however, is very complex and highly dynamic, with the

activation of Src and Rho GTPases on the one hand

and deactivation of ezrin and Rho GTPases on the

other. With PRL-3 being membrane bound and

PI(4,5)P2 being the highest abundant phosphoinositide

and a crucial part of the membrane in many respects

[92,96,97], this regulation essentially needs to be highly

dynamic and tightly regulated. Nevertheless, this inter-

action would destabilize focal adhesions and could

regulate focal adhesion turnover, leading to enhanced

motility and invasiveness. Further studies are necessary

to evaluate this hypothesis.

Expression and activity of matrix metalloproteinases

(MMP) is affected by PRLs. MMPs are extracellular

secreted proteins with a key function in tumour metasta-

sis [98]. Increased MMP2 (but not MMP9) activity and

expression levels have been found in PRL-3 stably trans-

fected LoVo cells [73]. PRL-3-induced invasion in these

cells was dependent on MMP2 upregulation and

ERK1 ⁄ 2 activation. PRL-3 also downregulated the

expression of the MMP2 inhibitor TIMP2, explaining,

at least in part, the activation of MMP2. Recently, Lee

et al. [99] investigated expression levels of several MMPs

in PRL-3-overexpressing colorectal DLD-1 cells.

MMP2 was also found to be enhanced, and MMP2

knockdown partially inhibited cell migration and inva-

sion. In addition, migration and invasion of DLD-1-

PRL3 cells was completely inhibited by small interfering

RNA (siRNA) knockdown of MMP7, whereas the over-

expression of MMP-7 increased migration. In agreement

with earlier studies, PRL-3 acted through oncogenic

pathways including PI3K ⁄Akt and ERK1 ⁄ 2 [99].A recent study revealed that intermediate-conduc-

tance Ca2+-activated K+ (KCNN4) channels were

upregulated in ectopically PRL-3 expressing LoVo

cells, and this upregulation was nuclear factor-jB(NF-jB)-dependent, revealing a novel pathway that

PRL-3 can interfere with. Blocking of KCNN4 chan-

nels inhibited PRL-3-induced cell proliferation and

arrested the cell cycle at the G2 ⁄M phase, indirectly

suggesting that PRL-3 facilitates G2 ⁄M transition in

this setting [100].

PRL-3 was also described to play a role in cell cycle

regulation in normal cells [84]. Tight control of PRL-3

basal expression in MEF cells appears to be important

to ensure cell cycle progression by facilitating G1 ⁄Stransition (as opposed to G2 ⁄M transition in LoVo

cancer cells). Its overexpression in MEF cells led to G1

arrest downstream of p53 via a PI3K-Akt-mediated

negative feedback loop, in which initial levels of PRL-3

activated the PI3K-Akt pathway but subsequent higher

levels of PRL-3 correlated with a decrease in activated



Akt. A decrease of PRL-3 expression levels also led to

cell cycle arrest through increased p53 expression and

via cyclin-dependent kinase 2 (CDK2), relying on an

intact p53 pathway. Interestingly, in the global study

by Ewing et al. [77], CDK2 was found to be a PRL-3

interacting protein.

These results appear to contradict the role of PRL-3

in cancer; however, Basak et al. [84] suggested that, as

a result of multiple mutations in cancer cells, particu-

larly in later (metastatic) stages, and with p53 loss of

function being a very common mutation, high expres-

sion levels of PRL-3 might not succeed in inducing cell

cycle arrest, and other functions of PRL-3 might pre-

vail. It is now important to dissect the primary role of

PRL-3 in healthy cells, whether it is related to cell

migration, cell cycle regulation or both, and whether

the activity in cancer is a malfunction or hyperactivity

of a normal function. Min et al. [30] addressed the

ability of PRL-3 to regulate p53 in cancer cells. In

agreement with the results obtained in MEF cells,

PRL-3 upregulation in HCT116 colorectal cancer cells

led to a decrease in p53 expression; however, it did not

lead to cell cycle arrest but to inhibition of p53-medi-

ated apoptosis [30]. An earlier, more detailed study on

PRL-1 investigated the mechanism of action of PRL-1

on p53 [101], and this is discussed below. PRL-3 was

described to act on p53 through the same mechanisms

involving mouse double minute 2 (MDM2) stabiliza-

tion via PI3K ⁄Akt signalling and also increased tran-

scription of protein with a RING-H2 domain

(PIRH2), both leading to p53 inactivation [30].

PRL-3 appears to play a role in drug resistance in

AML. PRL-3 was found at elevated levels in AML

patients and, in six out of nine patient samples, the

overexpression was correlated with internal tandem

repeat duplication of fms-like tyrosine kinase 3 (FLT3-

ITD), a mutation that occurs in approximately 25% of

AML patients. Zhou et al. [67] reported that, in AML

MOLM-14-cells, PRL-3 acts downstream of FLT3-

ITD through STAT5 and STAT3 (but not through

Akt) activation and upregulation of McI-1, which is

known to contribute to a resistance to chemotherapy

when it is highly abundant. In addition, PRL-3 was

shown to bind histone deacetylase 4 in MOLM-14 cell

lysate [67].

PRL-1


regulation

Initial studies reported that PRL-1 is expressed at high

levels in growing rat hepatic cells, rat intestinal epithe-

lia and some tumour cell lines, and also that it could

modulate cell growth or cell differentiation in a tissue-

dependent manner [17,25,102]. Expression of PRL-1

was found to be induced by the Egr-1 transcription

factor in liver regeneration and mitogen-activated cells

[103]. In normal adult human tissues, the PRL-1

mRNA expression pattern is widespread, although the

expression levels are variable in different tissues [104].

Endogenous PRL-1 was found to be expressed at

high levels in lymph node metastases of adenocarcino-

mas [105]. PRL-1 is also overexpressed in different

cancer cells (lung cancer, pancreatic cancer), conferring

increased cell motility and invasive properties that can

be counteracted when PRL-1 expression is knocked

down [14,106–108].

Although PRL-1 was first reported as a nuclear pro-

tein [17,102], it preferentially localizes (similar to the

other PRLs) in the plasma membrane and intracellular

membranes as a result of its farnesylation [10,23,25].

In mitotic cells, PRL-1 can localize to centrosomes

and the mitotic spindle in a farnesylation-independent

manner, colocalizing with a-tubulin (which physically

interacts with PRL-1 in vitro), and farnesylation defec-

tive mutants are reported to be associated with mitotic

defects [25].

Besides the interaction with a-tubulin, PRL-1 was

shown to interact with activated transcription factor

(ATF)-7 [109] (currently named ATF-5 ⁄ATF-X tran-

scription factor; a member of the ATF ⁄CREB family

of basic leucine zipper ⁄bZIP proteins). The interaction

involves the catalytic domain and a short adjacent

C-terminal region in PRL-1, as well as the bZIP

domain of ATF-7. ATF-7 was dephosphorylated

in vitro to some extent by PRL-1 and, to date, it is the

only proposed substrate for PRL-1. Recently, the

RhoA inhibitor p115 Rho-GTPase-activating protein

(RhoGAP) was described as a novel PRL-1 interacting

protein [110]. PRL-1 also interacts with different phos-

phoinositides (mainly mono- and di-phosphorylated)

and phosphatidic acid in vitro through the C-terminal

polybasic sequence, which cooperates with the farnesy-

lation to stabilize the protein at the membrane [10].

No phosphatase activity against phosphatidylinositol

phosphates was found for PRL-1 [10,111].

As noted above, another regulatory mechanism of

PRL-1 is oxidation. Endogenous PRL-1 in mammalian

retina cells and isolated retina tissue underwent revers-

ible inactivation by disulfide bond formation under

oxidative stress, and PRL-1 was reactivated by the glu-

tathione cellular redox system. Oxidative stress also

increased PRL-1 expression levels, suggesting that

PRL-1 can play additional roles in the oxidative stress

response [111].



Similar to PRL-3, PRL-1 is a p53 target. The PRL-

1-encoding gene contains a p53-binding element and

its mRNA transcription was reported to be regulated

by p53 [101].

Signalling pathways affected by PRL-1

Diverse focal adhesion components are regulated by

PRL-1. p130Cas phosphorylation and protein levels

were found to be downregulated in HeLa cells when

PRL-1 expression was knocked down or when the

PRL inhibitor thienopyridone was applied (this was

also the case for PRL-3) [112]. Another study reported

that the levels of Src and p130Cas were decreased

upon PRL-1 stable knockdown in A549 cells, whereas

no change in FAK expression was detected [106]. Nev-

ertheless, total tyrosine FAK phosphorylation and

Tyr397 phosphorylation levels were continuously ele-

vated when plating these cells in fibronectin, together

with a decrease in membrane protrusions and reduced

actin fiber extensions that could indicate decreased

adhesion turnover upon PRL-1 knockdown [106]. Fur-

thermore, ectopic overexpression of PRL-1 in HEK293

cells increased the autophosphorylation of Src and the

phosphorylation of FAK and p130Cas [108]. In addi-

tion, it was shown that the overexpression of PRL-1 in

A459 cells decreased the levels of vinculin, paxillin and

E-cadherin [14].

PRL-1 can modulate the activation of the small

GTPases RhoA, RhoC, Rac1 and Cdc42. In SW480

colon cancer cells, ectopic overexpression of PRL-1 led

to the activation of RhoA and RhoC, as well as the

inactivation of Rac1, and had no effect on Cdc42 [28].

Also, PRL-1-induced cell motility and invasion were

dependent on the effector Rho kinase (ROCK) in these

cells. As seen in SW480 cells, Nakashima and Lazo

[14] showed that the ectopic overexpression of PRL-1

in A549 lung cancer cells caused the activation of

RhoA (which depends on the PRL-1 catalytic activity)

and induced cell invasion and motility through Rho

kinase activation. PRL-1 overexpression in A549 cells

inactivated Rac1 and Cdc42, although this was inde-

pendent of PRL-1 catalytic activity, which suggested

that the PRL-1-promoted cell motility in A549 cells

was a result of RhoA activation and not dependent on

Rac1 and Cdc42 [14]. Interestingly, the stable knock-

down of PRL-1 also inactivated Rac1 and Cdc42 in

A549 cells (RhoA activation was not analyzed) but

only when the cells were plated in fibronectin [106].

These results reflect a complex and tight regulation of

the Rho proteins by PRL-1.

Recently, a mechanistic explanation about how

PRL-1 can activate ERK1 ⁄ 2 and RhoA signalling was

provided [110]. Through phage display screening, the

GTPase activating protein p115 RhoGAP was found

to be a new PRL-1 interacting partner. This interac-

tion involves a short motif within the p115 RhoGAP

SRC homology (SH) 3 domain. Interestingly, the crys-

tal structure of PRL-1 in complex with the p115 Rho-

GAP peptide identified in the screening revealed a

novel mode of interaction between the SH3 domain

and PRL-1 that is excluded from the canonical interac-

tion SH3 domain ⁄ ligand PxxP domain (which is absent

in PRL-1). This could be advantageous for further

drug development. It was observed that p115 RhoGAP

downregulates cell migration, ERK1 ⁄ 2 phosphoryla-

tion and RhoA activation in HEK293 cells, both in

PRL-1 stably transfected and in control cells. p115

RhoGAP also physically interacts with mitogen-acti-

vated protein ⁄ extracellular signal-regulated kinase

kinase kinase 1 (MEKK1) (and inhibits it) [113] and

RhoA [114]. However, when overexpressing PRL-1,

co-immunoprecipitation of p115 RhoGAP with

MEKK1 was greatly reduced, whereas ERK1 ⁄2 activa-

tion was enhanced. Furthermore, immunoprecipitated

p115 RhoGAP from PRL-1 overexpressing cells

showed lower GAP activity compared to control cells,

suggesting that PRL-1 can regulate p115RhoGAP

activity and thus RhoA activation. Furthermore, PRL-1

blocked the interaction between p115 RhoGAP and

RhoA. Thus, it appears that, through direct interaction

with p115 RhoGAP, PRL-1 plays a role in the modu-

lation of ERK1 ⁄2 and RhoA activation by sequester-

ing this negative regulator of MEKK1 and RhoA.

Whether the catalytic activity of PRL-1 has any influ-

ence in this process was not determined. Since the cat-

alytic activity is necessary for the PRL-induced cell

migration, as well as RhoA and ERK1 ⁄ 2 activation

[14,28,82], it would be interesting to investigate

whether this mode of action is based only on protein–

protein interactions.

The effects of PRL-1 on cell migration and invasion

can be partly mediated by an increased activity of

MMP2 and MMP9. Luo et al. [108] showed that

HEK293 cells stably overexpressing PRL-1 had ele-

vated levels (and activity) of MMP2 and MMP9. This

effect was mediated through the activation of Src by

increasing the phosphorylation of its Tyr416 (where

the residue number refers to chicken Src, correspond-

ing to Tyr419 in humans), leading to an increased

phosphorylation of p130Cas and FAK, and also

through the activation of ERK1 ⁄ 2. Moreover, PRL-1-

induced Src and ERK1 ⁄ 2 activation appear to control

the transcriptional upregulation of MMPs by activa-

tion of the transcription factors AP-1 and Sp-1. Simi-

lar regulation of MMPs, Src and ERK1 ⁄ 2 by PRL-1



was found in the lung cancer cell lines A549 and

H1299, where high levels of endogenous expression of

PRL-1 correlated with increased MMP2 and MMP9

expression levels and increased Src and ERK1 ⁄ 2 acti-

vity. Decreased cell migration and invasion was

observed when the expression of PRL-1 was knocked

down or when MMP or Src activity was inhibited in

these cell lines. In addition, the ectopic overexpression

of PRL-1 induced the expression of MMP2 and

MMP9 in H1299 cells, as observed in HEK293 cells

[108].

Similar to PRL-3, PRL-1 is implicated in cell cycle

regulation. The overexpression of PRL-1 in D27 ham-

ster pancreatic ductal epithelial cells induced cell cycle

progression, promoting entry into the S phase, upregu-

lating CDK2 activity and cyclin A protein levels, and

downregulating p21Cip1 ⁄ Waf1 levels [7]. The ectopic

expression of a catalytic defective mutant in HeLa cells

showed delayed progression through mitosis but no

other effects through the cell cycle [25].

Min et al. [101] reported that PRL-1 downregulates

p53 via a negative feedback mechanism. Endogenous

and exogenous p53 levels were reduced via ubiquitina-

tion when overexpressing PRL-1 in HCT116 and HeLa

cells. In addition, p53 levels were elevated when PRL-1

expression was suppressed by siRNA. When PRL-1

was overexpressed, an increased transcription of the

p53 ubiquitin ligase PIRH2 was observed, which was

mediated by the serum response factor target EGR1

(which, in turn, was transcriptionally activated by

PRL-1 overexpression). Furthermore, an increase in

Ser473 Akt phosphorylation was observed upon PRL-

1 overexpression leading to the phosphorylation of

MDM2, which can function both as a p53 ubiquitin

ligase and an inhibitor of p53 transcriptional activa-

tion. Both PRL-1-mediated p53 degradation pathways

were found to be independent. Thus, PRL-1 and PRL-3

might contribute to tumour development by the inhibi-

tion of p53-mediated apoptosis.

PRL-2


regulation

The human PRL-2-encoding gene was identified in the

BRCA1 locus of chromosome 1 [18]. Subsequently,

PRL-2 was also identified in mice, and northern blot

analysis of PRL-2 mRNA showed a preferential

expression in mouse skeletal muscle [19]. More

recently, by in situ hybridization, it was shown that

PRL-2 mRNA is almost ubiquitously expressed at high

levels in normal adult human tissues [104].

To date, the only reported PRL-2 interacting protein

is the b-subunit of Rab geranylgeranyltransferase II

(bGGT II) [5,24]. The geranylgeranyl transferase is a

heterodimeric enzyme composed of a and b subunits,

and incorporates C20 geranylgeranyl isoprenoids into

proteins containing a CAAX motif. The C-terminal

variable region of PRL-2 is required for the interaction

(which is specific for PRL-2 but not for PRL-1 or

PRL-3), and prenylation of PRL-2 is also necessary,

although PRL-2 is not a substrate of bGGT II [24].

This interaction was proposed to be a regulatory

mechanism of GGTII activity because the binding of

bGGT II to PRL-2 and to the aGGT II subunit is

mutually exclusive [24]. No physiological substrate has

yet been found for PRL-2.

Downregulation of the enzymatic activity by a disul-

fide bond between Cys49 and Cys104 has been demon-

strated for PRL-1 and for PRL-3. The same would be

expected for PRL-2, although this remains to be

addressed.

PRL-2 in cancer and signal transduction

Among the PRL group of proteins, PRL-2 is the least

studied member. In particular, its role in cancer has

not been addressed in depth, even though it was ini-

tially reported that the ectopic expression of PRL-2 is

involved in cell transformation and tumour progres-

sion [6]. Two other studies showed the expression of

PRL-2 in different primary and metastatic tumours

[105,115], but detailed studies about the regulation of

signalling mechanisms and PRL-2 in cancer biology

were (and are) still missing. In recent years, a number

of studies reported that PRL-2 is also overexpressed in

different cancer cell lines and ⁄or tumour samples (pan-

creatic, breast and lung cancer) and, more importantly,

it was shown that PRL-2 is associated with tumour

progression [4,5,107]. Also, an effect on malignant pro-

gression and metastasis by ectopic overexpression of

PRL-2 in hematopoietic cells was reported [116].

Taken together, these recent findings demonstrate that

PRL-2, similar to PRL-1 and PRL-3, should be con-

sidered as an oncogenic protein, and emphasize the

importance of carrying out individual studies of the

three members of this group of phosphatases.

Stephens et al. [107] found that PRL-1 and PRL-2

(but not PRL-3) are overexpressed in pancreatic cancer

cell lines and pancreatic tumours. Significant results in

decreased cell growth, cell migration and soft colony

agar formation were observed only when performing

double knockdown of PRL-1 and PRL-2 expression in

PANC1 and MIA PaCa-2 pancreatic cancer cell lines,

suggesting the overlapping functions of both proteins.



These effects can be mediated by PI3K and Erk1 ⁄ 2signalling because there was a decrease in Akt

serum-induced phosphorylation in both cell lines and

decreased Erk1 ⁄ 2 activation in MIA PaCa-2, whereas

this was increased in PANC1 cells [107].

PRL-2 mRNA levels are elevated and associated with

prognosis in pediatric AML [117]. Akiyama et al. [116]

showed that, when stably overexpressing Flag-PRL-2

into the murine pre-B cell line BaF3ER, different malig-

nant features are observed (including increased cell

migration). These cells displayed increased erytropoie-

tin and interleukin-3 dependent cell growth and, when

stimulated with erytropoietin or interleukin-3, the phos-

phorylation levels of STAT5 were two- or five-fold

higher, respectively, than the stimulated control-trans-

fected cells. Also, PRL-2 enhanced erytropoietin-

induced cell growth in mouse primary bone marrow

transduced cells. Thus, a contribution of PRL-2 in

hematopoietic malignancies was suggested, and this

may involve STAT5 mediated signalling. Because the

injection of PRL-2 overexpressing cells did not result in

tumours in nude mice, and also because PRL-2 overex-

pressing cells were still dependent on proliferation med-

iated by growth factors, PRL-2 may need additional

oncogenic factors to achieve a complete malignant phe-

notype in hematopoietic cells [116].

It was previously shown that the PRL mRNAs were

elevated in different breast cancer cell lines, although

significant differences in elevated mRNA levels

between neoplastic and normal tissue were only found

for PRL-3 [118]. Hardy et al. [5] observed that mRNA

levels of PRL-2 were elevated in primary breast

tumours compared to normal tissue levels in 16 out of

19 patients. More remarkably, PRL-2 was greatly

overexpressed in metastatic lymph nodes compared to

primary tumours. Hardy et al. [5] demonstrated that

PRL-2 plays a role in cell migration and the transfor-

mation process in different breast cancer cells. Ectopic

expression of PRL-2 in fully transformed TM15 and

DB7 cell lines increased colony formation in soft agar

and cell migration. Knockdown of PRL-2 in MDA-

MB-231 cells decreased anchorage-independent growth

and cell migration. When implanting cells overexpress-

ing PRL-2 into the mouse mammary fat pad, an

increase in tumour size and weight was observed com-

pared to control animals (which was also correlated

with increased ERK1 ⁄ 2 phosphorylation). On the

other hand, an effect in mice breast tumour generation

only took place in PRL-2 transgenic mice against an

oncogenic ErbB2 background (which exhibited acceler-

ated tumour development, increased ERK1 ⁄ 2 phos-

phorlyation and had no effect in Akt activation) [5].

Similar to the reported findings in hematopoietic can-

cer cells [116], PRL-2 alone would not be sufficient to

trigger oncogenesis.

Wang and Lazo [4] reported that PRL-2 is involved

in lung cancer cell migration and invasion through the

ERK1 ⁄ 2 signalling pathway. It was found that PRL-2

was overexpressed in four lung cancer cell lines com-

pared to the CCL202 normal fibroblast lung cell line.

When knocking down the PRL-2 expression with

siRNA, migration and invasion of A549 cells was inhib-

ited. Decreased levels of p130Cas, previously described

in Hela cells [112], and vinculin were found, whereas

paxilin levels remained unchanged upon PRL-2 knock-

down. Silenced expression of PRL-2 did not have any

effect on Src protein levels or phosphorylation status

(neither in Akt or p53) in A549 cells, but it led to a

decrease in ERK1 ⁄2 phosphorylation. In addition, ecto-

pic overexpression of PRL-2 induced cell migration, cell

invasion and Erk1 ⁄ 2 phosphorylation (and its nuclear

translocation), and both the catalytic activity and farn-

esylation were necessary. As a result of their findings

regarding Src kinase, Wang and Lazo [4] suggested that

PRL-2, compared to PRL-3 and PRL-1, signals

through different mechanisms in A549 cells (see below).

Nevertheless, Akt activation was downregulated by

PRL-2 knockdown in pancreatic cancer cell lines [107],

which reflects that the actions of PRL-2 likely depend

on the molecular context in different cancers.

Finally, similar to PRL-1, PRL-2 is involved in cell

cycle regulation by promoting the G1 to S transition

through the downregulation of p21Cip1 ⁄ Waf1 [7].

Differences in molecular mechanismsof the PRLs

Signalling pathways

Some common signalling mechanisms are shared by

the three PRLs, such as the activation of ERK1 ⁄2or the regulation of the focal adhesion contacts via

p130Cas (Fig. 4). However, some differences in focal

adhesion contact regulation by the PRLs can be

found (e.g. at the Src kinase level). Src kinase

activity is regulated by autophosphorylation at its

Tyr419 residue (meaning activation) or by phosphor-

ylation at its C-terminal Tyr530 residue (meaning

inactivation) (where residue numbering refers to the

human sequence) by Csk [119]. PRL-3 downregulates

Csk and thereby activates Src, but this mechanism has

not been studied for PRL-1 or PRL-2. However, a dif-

ferent mechanism of Src activation by PRL-1 was

observed through the increase of tyrosine phosphoryla-

tion at the Src Tyr419 residue, which was not observed

for PRL-3. Also, the protein levels of Src were



downregulated upon PRL-1 knockdown. By contrast,

PRL-2 knockdown decreased neither the protein, nor

the phosphorylation levels of Src, although p130Cas

levels were diminished. Therefore, PRL-2 was proposed

to use a Src-independent mechanism of p130Cas signal-

ling. Further studies are needed to completely under-

stand this observation.

Differences in the regulation of Rho proteins by PRL-

1 and PRL-3 are found (the effects of PRL-2 in the Rho

GTPase family have not yet been analyzed). Both acti-

vate RhoA and RhoC; however, PRL-3 can also down-

regulate RhoA activity. PRL-3 downregulates Rac1

and, to date, an effect on Cdc42 has not been observed.

The effects of PRL-1 on Rac1 and Cdc42 are not yet

completely understood. Different modes of regulation

can be attributed to distinct cellular contexts or to the

dynamic regulation of Rho proteins during cell adhesion

and migration. Whether PRL-3 could also interact with

p115 RhoGAP (as is the case for PRL-1) and regulate

the activities of RhoA and ERK1 ⁄ 2 in this way has not

been addressed yet.

The MMPs are positively regulated by PRL-1 (via

Src ⁄ERK1 ⁄ 2 and increasing MMP2 and MMP9

expression) and by PRL-3 (via integrin b1 or

Fig. 4. Overview depicting the current knowledge of the signalling pathways affected by the PRL phosphatases and the outcome in cell

migration and proliferation. Arrows indicate positive regulation; crossed lines indicate negative regulation; and question marks indicate either

not yet understood or not studied processes. Detailed explanations are provided in the text.



PI3K ⁄Akt and ERK1 ⁄2 and increasing MMP2 and

MMP7; but not MMP9 activity and expression).

Further investigations are needed to understand which

molecular mechanisms of MMP activation are shared

by PRL-3 and PRL-1 and which are not, and no stud-

ies are available for PRL-2 in this respect.

It appears that PRL-1 and PRL-3 share a common

mechanism of p53 downregulation through the activa-

tion of the ubiquitin ligases MDM2 and PIRH2. In

addition, both PRL-1 and PRL-3 are p53 targets. PRL-

1 downregulates the cyclin dependent kinase inhibitor

p21; PRL-2 does this as well, however, there are no stud-

ies addressing the putative regulation of p53 by PRL-2.

Furthermore, the activation of the EMT has only

been studied for PRL-3 and not yet for PRL-1 or

PRL-2.

The discovery of the physiological substrates of the

PRL phosphatases and the correlation with their cellular

phenotypes is probably one of the most important ques-

tions that still remains unanswered. As noted above, the

only putative substrate identified for PRL-1 is the tran-

scription factor ATF-7 ⁄ATF-5. Subsequently, no fur-

ther studies have been carried out aiming to establish

whether ATF-7 ⁄ 5 is a bona fide substrate and to under-

stand its physiological relevance. Interestingly, both

PRL-2 and ATF-5 mRNAs were found to be overex-

pressed in the L1236 Hodgkin’s lymphoma cell line

[120]. It would be interesting to determine whether

PRL-2 is also an interacting partner of ATF-5. ATF-5 is

widely expressed in human carcinomas [121] and regu-

lates cell differentiation, cell survival and apoptosis

[122]. In glioblastoma cells, ATF-5 ⁄ 7 loss of function

lead to apoptosis [123] and the prosurvival protein

BCL-2 is a downstream target of ATF-7 [124]. Whether

phosphorylation plays a role in the regulation of ATF-

7 ⁄ATF-5, and whether it could be affected by PRL-1 (or

other PRLs), requires future studies.

Of the putative PRL-3 substrates, only phosphoinosi-

tides have been tested as substrates for PRL-1.

Strikingly, PRL-1 does not show activity against phos-

phoinositides [10,111] (V. McParland and M. Kohn,

unpublished observations), whereas PRL-3 dephosph-

orylates PI(4,5)P2 [50]. This finding could indicate,

despite sequence and structural similarities, that the

PRL phosphatases possess important differences in

function and that the presence of only a very few dissim-

ilarities in sequence and structure could make a big dif-

ference with respect to substrate specificities.

Comparison of PRL structures and sequences

What could those dissimilarities in sequence and

structure be? Figure 5 shows the NMR structures of

PRL-3 [32,35] and the X-ray crystal structures of

PRL-1 [31,34]. The different methods by which these

structures were obtained should be kept in mind

when comparing these structures because differ-

ences can occur due to the different methods

employed.

The PRL-3 structures were solved in the apo form

and, in both structures, PRL-3 is shown in the reduced

state with respect to Cys49 and Cys104. Overlaying

both PRL-3 structures shows that the WPFDD loop is

very flexible but, in both structures, the loop is not

closing the active site, suggesting that PRL-3 is in an

inactive but reduced state and potentially ready to

accept substrates (Fig. 5A). PRL-1 was crystallized in

the apo form and bound to a sulfate ion. By contrast

to PRL-3, all apo structures contain a disulfide bond

between Cys49 and Cys104, showing the enzyme in its

inactive, oxidized state (Fig. 5B). Nevertheless, the apo

structures align well with the sulfate ion bound

structures, and the flexible WPFDD loop in all cases is

in its closed form (Fig. 5C). Compared to PRL-1, and

possibly as a result of the lack of the disulfide bond,

the active site p-loop of PRL-3 adopts a different con-

formation (see Arg110) and is flatter in these structures

(Fig. 5C) [34].

X-ray structures are snapshots of proteins. However,

it is curious that all three apo structures of PRL-1

show no difference in WPFDD loop conformations or

the oxidation state. Possibly, disulfide bond formation

induces a conformational change by an unknown

mechanism, which closes the protein active site, pre-

venting substrates from binding. The preference for

PRL-1 being oxidized, whereas PRL-3 is not, might

hint at the redox potential of PRL-1 being different

from that of PRL-3.

The amino acids in the active sites of PRL-1 and

PRL-3 are completely conserved; all nonconserved res-

idues are at other sites in the proteins [34]. Differing

amino acids closest to the active site comprise Ile141

and Pro77Gly78Lys79 in PRL-3, which correspond to

Phe141 and Ser77Asn78Gln in PRL-1 (Fig. 1). It was

proposed that the difference in the three amino acids

77–79 could lead to a higher flexibility in the PRL-3

compared to the PRL-1 WPFDD loop [32,34]. In

addition, amino acids 77 and 78 in PRL-3 introduce a

higher hydrophobicity than in PRL-1 at this stretch,

and the additional proline induces conformational

restraints. Similarly, Ile141 in PRL-3 appears to be

flexible and solvent-exposed, whereas the correspond-

ing Phe141 in PRL-1 is buried in a helix in all

structures (Fig. 5C). Ile141 forms a network of

aliphatic site chains with Ile130 and Leu146, which

would not only result in a difference in the position of



the involved helices, but also add an additional

hydrophobic interface to the surface of PRL-3. We

observed that PRL-3 prefers phosphoinositides with

long lipid chains (M. Bru and M. Kohn, unpublished

results) and, interestingly, the lipid chains are able to

reach the hydrophobic stretches (as seen from mole-

cular docking experiments, X. Li and M. Kohn,

unpublished results). Thus, in addition to the different

conformations of the p-loop and WPFDD-loop, these

hydrophobic stretches could play a role with respect to

the difference in substrate (particularly phosphoinosi-

tide) recognition by PRL-3 and PRL-1, as also previ-

ously suggested for Ile141 [32].

Conclusions

In conclusion, notwithstanding the progress made in

understanding PRL molecular mechanisms, many

questions remain unanswered. Future studies are

needed to elucidate the physiological substrates of the

PRL family. It will be important to determine whether

substrates are shared by the three PRLs or if they act

only on different substrates. Since the PRLs appear to

be very similar but show distinct differences (e.g. in

substrate specificity and expression patterns), studies

carried out under ectopic overexpression conditions

need to be considered with caution when comparing

the PRLs, and cell types need to be chosen (particu-

larly in healthy cells) according to their relevance

in vivo. Together, this will help to explain the regula-

tion and function of the PRLs not only under physio-

logical conditions, but also in the context of tumours,

and could help in the development of different thera-

peutic strategies [2].

Acknowledgements

This work was supported by the German Science

Foundation (Deutsche Forschungsgemeinschaft, DFG)

within the Emmy-Noether program for M.K., and by

the EMBL and Marie Curie Action EMBL Interdisci-

plinary Postdoc fellowships for X.L. and P.R.

Fig. 5. Structural comparison between

PRL-1 and PRL-3. (A) PRL-3 apo structures

(1V3A, white; 1R6H-model01, green). (B)

PRL-1 apo structures (1RXD, green; 1X24,

yellow; 1ZCK, magenta). (C) Overlay of

PRL-1 (1XM2, white; 1RXD, cyan; 1X24,

yellow; 1ZCK, pink) and PRL-3 (1V3A,

magenta). The sulfate ion is from 1XM2.

Amino acids are numbered according to

protein database files.



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molecular mechanisms of the prl phosphatases

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