• cancer;
  • cell signalling;
  • dual specificity phosphatases;
  • metastasis;
  • phosphatase of regenerating liver;
  • protein phosphatases


  1. Top of page
  2. Abstract
  3. Common features of the PRLs
  4. Comparison of structural features of the PRLs with other related phosphatases
  5. PRL-3
  6. PRL-1
  7. PRL-2
  8. Differences in molecular mechanisms of the PRLs
  9. Conclusions
  10. Acknowledgements
  11. References

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 phosphatases of regenerating liver, compare them to related phosphatases, and review recent reports about each phosphatase. Finally, we discuss similarities and differences between the phosphatases of regenerating liver, focusing on their molecular mechanisms and signalling pathways.


acute myeloid leukaemia


activated transcription factor


β-subunit of Rab geranylgeranyltransferase II


cell division cycle 14 homologue


cyclin-dependent kinase 2


C-terminal Src kinase


elongation factor 2


epithelial–mesenchymal transition


extracellular signal-regulated kinase


focal adhesion kinase


FK506-binding protein 38


guanosine triphosphate phosphohydrolase


kinase-associated phosphatase


mouse double minute 2


mouse embryonic fibroblast


myocyte enhancer factor 2C


mitogen-activated protein/extracellular signal-regulated kinase kinase kinase 1


matrix metalloproteinase


ortho-methylfluorescein phosphate


polyC-RNA-binding protein 1


phosphatidylinositol 3-kinase


phosphatidylinositol 4,5-bisphosphate


protein with a RING-H2 domain


para-nitrophenyl phosphate


phosphatase of regenerating liver


phosphatase and tensin homologue


protein tyrosine phosphatase


protein tyrosine phosphatase mitochondrial 1


Rho-GTPase-activating protein


SRC homology 3 domain


small interfering RNA




transforming growth factor


vascular endothelial growth factor


vaccinia H1-related phosphatase

Common features of the PRLs

  1. Top of page
  2. Abstract
  3. Common features of the PRLs
  4. Comparison of structural features of the PRLs with other related phosphatases
  5. PRL-3
  6. PRL-1
  7. PRL-2
  8. Differences in molecular mechanisms of the PRLs
  9. Conclusions
  10. Acknowledgements
  11. References

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 development of cancer [2–5]. The PRLs promote cell proliferation, 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 cysteine that is prenylated, A is an aliphatic amino acid and X is any amino acid) [6]. In addition, the latter 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).


Figure 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 ( and the alignment was computed with clustalw_3d. The alignment was manually adjusted according to the superimposed structures.

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PRLs are classified into the family of dual specificity phosphatases (DSP) (also called vaccinia H1-like phosphatases) [21], which is a subgroup of the class I protein tyrosine phosphatase family defined by the conserved active site p-loop sequence HC(X)5R[S/T] [22]. PRLs are relatively small proteins of approximately 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 geranylgeranylated 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 catalytic 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 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 methods, although it could also mean that there are differences within the PRL family with respect to the ability/tendency to form oligomers. PRL-1 oligomerization, 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 farnesylation-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 phosphatase and tensin homologue (PTEN) [37]. It was speculated that this redox-mechanism not only regulates the activity of the phosphatases, but also could protect the catalytic Cys104 from further and irreversible oxidation [31,32]. Skinner et al. [38] reported that the reduction 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 conformational 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 regulate PRL-1 subcellular localization, but also the cellular 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, focusing 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 recognition mechanism is provided.

Comparison of structural features of the PRLs with other related phosphatases

  1. Top of page
  2. Abstract
  3. Common features of the PRLs
  4. Comparison of structural features of the PRLs with other related phosphatases
  5. PRL-3
  6. PRL-1
  7. PRL-2
  8. Differences in molecular mechanisms of the PRLs
  9. Conclusions
  10. Acknowledgements
  11. References

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 structurally most closely-related phosphatases [PTEN, vaccinia H1-related phosphatase (VHR), cell division cycle 14 homologue (CDC14), kinase-associated phosphatase (KAP)], PRLs lack helices and loops that can be important for substrate recognition [31,34]. In addition, all of those phosphatases, which are structurally most closely related, have very different substrate specificities, 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 residues similar to KAP and CDC14B. Nevertheless, it was proposed that significant structural rearrangements will likely occur upon association of PRL phosphatases 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.


Figure 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.

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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 phosphatase [51]. Replacement of alanine with serine enhanced the catalytic activity towards unnatural substrates 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 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 phosphatase 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 conservation 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 myotubularin 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 cysteine 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 mutation in PRL-3 did not lead to a change in activity against the unnatural substrate OMFP [32]. Considering that an alanine mutation (C49A in PRL-3) introduces a much less drastic change in electrostatic and steric properties than a tyrosine mutation (C71Y in PTEN) does, and that activities toward unnatural substrates 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 positioning of amino acids that are involved in substrate interactions or the catalytic mechanism. This idea is fueled by another interesting consideration: PTPMT1 contains the catalytically relevant ‘EEYE’ loop, in which Glu73 and Glu76 were shown to be essential for catalytic 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/or Glu76 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).


Figure 3.  Structural alignment of the putative ‘CXnE’ motif. The Glu residues adjacent to the regulatory cysteines in the crystal structures 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.

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  1. Top of page
  2. Abstract
  3. Common features of the PRLs
  4. Comparison of structural features of the PRLs with other related phosphatases
  5. PRL-3
  6. PRL-1
  7. PRL-2
  8. Differences in molecular mechanisms of the PRLs
  9. Conclusions
  10. Acknowledgements
  11. References

PRL-3 expression, interacting proteins and regulation

PRL-3 mRNA was found predominantly in the skeletal 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 development, and not in the human adult organism, as demonstrated at the mRNA and protein levels [13]. This finding could have important implications for any potential drug discovery against PRL-3 because inhibition of PRL-3 in the adult heart could lead to cardiotoxic 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 differentiated 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 prognoses and, particularly for colon cancer, high levels of PRL-3 were shown to be predictive for the development 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 malignancies, 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 β1 [71], all four of which have been reviewed [2], as well as stathmin [72] and nucleolin [15]. Recently, we described phosphatidylinositol(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 differences 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 phosphate [50]. Of the putative substrates, direct dephosphorylation 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 β1 is now considered to be indirectly affected by PRL-3 [2,73]. In independent experiments, however, the influence 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 partners have been identified: integrin α1 [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 α1, CDH22] or to the cytoskeleton [keratin 8, stathmin]. Noteworthy, nucleolin is localized to the cytoplasm and nucleus and is involved in cell proliferation [15] and FKBP38 is a cytosolic protein that regulates 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 considered 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 connection to another substrate, ezrin. Ezrin forms part of the ERM (ezrin–radixin–moesin) complex, which connects 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 prevention of ezrin binding to the plasma membrane [79]. On the other hand, PRL-3 has been reported to upregulate 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 submembraneous 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, depending on the functional context. This shows that the activity of PRL-3 takes place in a very complex environment, which in itself remains incompletely understood.

PRL-3 is subject to complex regulatory mechanisms. It is known that PRL-3 mRNA levels do not necessarily 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 hepatocellular carcinoma) cells [30,84], and ectopic expression 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 factors [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 vessel density [12], demonstrating that the expression of PRL-3 facilitates angiogenesis [2].

Snail is a transcription factor involved in the epithelial–mesenchymal transition (EMT). EMT is an important 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)β signalling in colon cancer metastasis. TGFβ signalling suppresses the metastasis of colon cancer cells potentially by inducing stress-induced apoptosis. It was demonstrated that TGFβ signalling inhibited the expression of PRL-3 in a mothers against decapentaplegic homologue (Smad) 3-dependent manner. Because a loss of TGFβ 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 suppressor 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 mesenchymal markers and downregulates epithelial markers, it was shown that PRL-3 promotes EMT [75,88]. It promotes 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 levels [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 activation 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)-3β [75,88]. Furthermore, 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 turnover 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 promote cell motility via many different pathways involving 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 α1 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 β1 in LoVo colon cancer cells leading to extracellular signal-regulated 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 regulator 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 activated 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 demonstrate that the Rho family of GTPases act downstream of PRL-3, and also show that the regulation is complex. 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 interaction 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 metastasis [98]. Increased MMP2 (but not MMP9) activity and expression levels have been found in PRL-3 stably transfected 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 invasion. In addition, migration and invasion of DLD-1-PRL3 cells was completely inhibited by small interfering RNA (siRNA) knockdown of MMP7, whereas the overexpression 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-conductance Ca2+-activated K+ (KCNN4) channels were upregulated in ectopically PRL-3 expressing LoVo cells, and this upregulation was nuclear factor-κB (NF-κB)-dependent, revealing a novel pathway that PRL-3 can interfere with. Blocking of KCNN4 channels 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/S transition (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, particularly in later (metastatic) stages, and with p53 loss of function being a very common mutation, high expression levels of PRL-3 might not succeed in inducing cell cycle arrest, and other functions of PRL-3 might prevail. 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-mediated 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) stabilization via PI3K/Akt signalling and also increased transcription 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].


  1. Top of page
  2. Abstract
  3. Common features of the PRLs
  4. Comparison of structural features of the PRLs with other related phosphatases
  5. PRL-3
  6. PRL-1
  7. PRL-2
  8. Differences in molecular mechanisms of the PRLs
  9. Conclusions
  10. Acknowledgements
  11. References

PRL-1 expression, interacting proteins and regulation

Initial studies reported that PRL-1 is expressed at high levels in growing rat hepatic cells, rat intestinal epithelia 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 adenocarcinomas [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 protein [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 α-tubulin (which physically interacts with PRL-1 in vitro), and farnesylation defective mutants are reported to be associated with mitotic defects [25].

Besides the interaction with α-tubulin, PRL-1 was shown to interact with activated transcription factor (ATF)-7 [109] (currently named ATF-5/ATF-X transcription 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 phosphoinositides (mainly mono- and di-phosphorylated) and phosphatidic acid in vitro through the C-terminal polybasic sequence, which cooperates with the farnesylation 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 reversible inactivation by disulfide bond formation under oxidative stress, and PRL-1 was reactivated by the glutathione 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]. Nevertheless, total tyrosine FAK phosphorylation and Tyr397 phosphorylation levels were continuously elevated 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]. Furthermore, ectopic overexpression of PRL-1 in HEK293 cells increased the autophosphorylation of Src and the phosphorylation of FAK and p130Cas [108]. In addition, 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 independent 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 knockdown 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 interaction involves a short motif within the p115 RhoGAP SRC homology (SH) 3 domain. Interestingly, the crystal structure of PRL-1 in complex with the p115 RhoGAP peptide identified in the screening revealed a novel mode of interaction between the SH3 domain and PRL-1 that is excluded from the canonical interaction 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 phosphorylation and RhoA activation in HEK293 cells, both in PRL-1 stably transfected and in control cells. p115 RhoGAP also physically interacts with mitogen-activated 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 activation 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 modulation of ERK1/2 and RhoA activation by sequestering this negative regulator of MEKK1 and RhoA. Whether the catalytic activity of PRL-1 has any influence in this process was not determined. Since the catalytic 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 elevated 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, corresponding 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 activation of the transcription factors AP-1 and Sp-1. Similar 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 activity. 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 hamster pancreatic ductal epithelial cells induced cell cycle progression, promoting entry into the S phase, upregulating 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 ubiquitination 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 activation. 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 inhibition of p53-mediated apoptosis.


  1. Top of page
  2. Abstract
  3. Common features of the PRLs
  4. Comparison of structural features of the PRLs with other related phosphatases
  5. PRL-3
  6. PRL-1
  7. PRL-2
  8. Differences in molecular mechanisms of the PRLs
  9. Conclusions
  10. Acknowledgements
  11. References

PRL-2 expression, interacting proteins and 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 β-subunit of Rab geranylgeranyltransferase II (βGGT II) [5,24]. The geranylgeranyl transferase is a heterodimeric enzyme composed of α and β 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 βGGT II [24]. This interaction was proposed to be a regulatory mechanism of GGTII activity because the binding of βGGT II to PRL-2 and to the αGGT II subunit is mutually exclusive [24]. No physiological substrate has yet been found for PRL-2.

Downregulation of the enzymatic activity by a disulfide bond between Cys49 and Cys104 has been demonstrated 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 initially reported that the ectopic expression of PRL-2 is involved in cell transformation and tumour progression [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 (pancreatic, 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 progression 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 considered 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 Erk1/2 signalling 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 malignant features are observed (including increased cell migration). These cells displayed increased erytropoietin and interleukin-3 dependent cell growth and, when stimulated with erytropoietin or interleukin-3, the phosphorylation levels of STAT5 were two- or five-fold higher, respectively, than the stimulated control-transfected 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 overexpressing cells were still dependent on proliferation mediated by growth factors, PRL-2 may need additional oncogenic factors to achieve a complete malignant phenotype 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 transformation 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 overexpressing PRL-2 into the mouse mammary fat pad, an increase in tumour size and weight was observed compared 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 accelerated tumour development, increased ERK1/2 phosphorlyation and had no effect in Akt activation) [5]. Similar to the reported findings in hematopoietic cancer 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 compared to the CCL202 normal fibroblast lung cell line. When knocking down the PRL-2 expression with siRNA, migration and invasion of A549 cells was inhibited. Decreased levels of p130Cas, previously described in Hela cells [112], and vinculin were found, whereas paxilin levels remained unchanged upon PRL-2 knockdown. 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, ectopic overexpression of PRL-2 induced cell migration, cell invasion and Erk1/2 phosphorylation (and its nuclear translocation), and both the catalytic activity and farnesylation 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 mechanisms of the PRLs

  1. Top of page
  2. Abstract
  3. Common features of the PRLs
  4. Comparison of structural features of the PRLs with other related phosphatases
  5. PRL-3
  6. PRL-1
  7. PRL-2
  8. Differences in molecular mechanisms of the PRLs
  9. Conclusions
  10. Acknowledgements
  11. References

Signalling pathways

Some common signalling mechanisms are shared by the three PRLs, such as the activation of ERK1/2 or 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 phosphorylation 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 different mechanism of Src activation by PRL-1 was observed through the increase of tyrosine phosphorylation 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 signalling. Further studies are needed to completely understand this observation.


Figure 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.

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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 activate RhoA and RhoC; however, PRL-3 can also downregulate 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 β1 or 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 studies 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 activation 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 studies 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 questions that still remains unanswered. As noted above, the only putative substrate identified for PRL-1 is the transcription factor ATF-7/ATF-5. Subsequently, no further studies have been carried out aiming to establish whether ATF-7/5 is a bona fide substrate and to understand its physiological relevance. Interestingly, both PRL-2 and ATF-5 mRNAs were found to be overexpressed 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 regulates 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 phosphoinositides have been tested as substrates for PRL-1. Strikingly, PRL-1 does not show activity against phosphoinositides [10,111] (V. McParland and M. Köhn, unpublished observations), whereas PRL-3 dephosphorylates 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 dissimilarities in sequence and structure could make a big difference 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 differences can occur due to the different methods employed.


Figure 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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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 conformation (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, preventing 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 residues 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 corresponding 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. Köhn, unpublished results) and, interestingly, the lipid chains are able to reach the hydrophobic stretches (as seen from molecular docking experiments, X. Li and M. Köhn, 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 phosphoinositide) recognition by PRL-3 and PRL-1, as also previously suggested for Ile141 [32].


  1. Top of page
  2. Abstract
  3. Common features of the PRLs
  4. Comparison of structural features of the PRLs with other related phosphatases
  5. PRL-3
  6. PRL-1
  7. PRL-2
  8. Differences in molecular mechanisms of the PRLs
  9. Conclusions
  10. Acknowledgements
  11. References

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 (particularly in healthy cells) according to their relevance in vivo. Together, this will help to explain the regulation and function of the PRLs not only under physiological conditions, but also in the context of tumours, and could help in the development of different therapeutic strategies [2].


  1. Top of page
  2. Abstract
  3. Common features of the PRLs
  4. Comparison of structural features of the PRLs with other related phosphatases
  5. PRL-3
  6. PRL-1
  7. PRL-2
  8. Differences in molecular mechanisms of the PRLs
  9. Conclusions
  10. Acknowledgements
  11. References

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


  1. Top of page
  2. Abstract
  3. Common features of the PRLs
  4. Comparison of structural features of the PRLs with other related phosphatases
  5. PRL-3
  6. PRL-1
  7. PRL-2
  8. Differences in molecular mechanisms of the PRLs
  9. Conclusions
  10. Acknowledgements
  11. References
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