modulate the poly(A) tail length, presumably by deadeny- lation, and identify a broad role of deadenylation regulation in gene expression (Beilharz and Preiss 2007). For example, the six members of the Puf protein family bind to speciﬁc- sequence 39 UTR elements and regulate 10% of the yeast transcriptome (Olivas and Parker 2000; Gerber et al. 2004; Yosefzon et al. 2011). In yeast, the Puf1, Puf3, Puf4, and Puf5 proteins have all been shown to promote deadenyla- tion and degradation of speciﬁc subsets of yeast mRNAs (Olivas and Parker 2000; Tadauchi et al. 2001; Hook et al. 2007; Ulbricht and Olivas 2008). Mechanistic studies have demonstrated that Puf5 promotes deadenylation at least in part by direct interaction with Pop2 and thereby recruitment of the Ccr4 deadenylase (Goldstrohm et al. 2006, 2007), although Puf proteins may also recruit the deadenylase com- plexes through other interactions. In addition, Puf proteins can also repress translation independently of deadenylation and therefore might also promote deadenylation indirectly (Chritton and Wickens 2011). Other sequence-speciﬁc regulators of deadenylation in- clude the Vts1 protein, which binds to a subset of yeast mRNAs through a speciﬁc stem-loop structure (Aviv et al. 2006) and recruits the Ccr4/Pop2 deadenylase (Rendl et al. 2008). Similarly, the Cth1 and Cth2 proteins are zinc-ﬁnger RNA-binding proteins that regulate the deadenylation of a subset of mRNAs, perhaps through interactions with Dhh1 that interacts with Pop2 (Puig et al. 2005; Pedro-Segura et al. 2008). One anticipates that a growing set of mRNA- speciﬁc binding proteins will regulate deadenylation either by direct recruitment of the deadenylase complexes or by inhibiting translation initiation and thereby indirectly pro- moting deadenylation. Environmental control of deadenylation: Deadenylation is also regulated on a global scale in response to environmen- tal cues. For example, a variety of different stresses lead to a general inhibition of both Ccr4 and Pan2 deadenylation (Hilgers et al. 2006). Similar results occur in mammalian cells, suggesting that inhibition of deadenylation is a con- served aspect of the stress response (Gowrishankar et al. 2005, 2006). Inhibition of deadenylation during stress does not seem to require mRNAs to assemble in stress granules or P-bodies (see below), since deadenylation is still inhibited by stress in the presence of cyclohexmide (Hilgers et al. 2006), which prevents the formation of stress granules and P-bodies (Sheth and Parker 2003; Buchan et al. 2008). Dead- enylation is also inhibited during stress. Since the stress re- sponse often leads to a global decrease in translation initiation, a general inhibition of deadenylation might be re- quired to maintain a stable population of mRNAs. Deadeny- lation can also be reduced for some mRNAs when Hsp70 function is altered, which might mimic a stress response, al- though the basis or generality of this effect has not been de- termined (Duttagupta et al. 2003). Normal rates of deadenylation also appear to be de- pendent on the activity of the Pkh1 and Pkh2 kinases, which 676 R. Parker are activated by sphingolipids (Luo et al. 2011). This sug- gests that aspects of mRNA metabolism are modulated in response to lipid signaling. This interpretation is also sup- ported by the observation that, during heat stress, the for- mation of P-bodies, which are cytoplasmic mRNP aggregates of untranslating mRNAs, in conjunction with the mRNA decapping machinery (see below), requires sphingolipid synthesis, and exogenous phytosphingosine can stimulate P-body formation (Cowart et al. 2010). Interestingly, the effect of Pkh1 and Pkh2 on deadenylation rates is observed only in synthetic media, suggesting that this regulation is an integrative readout of both lipid and nutrient availability (Luo et al. 2011). An important area of future work will be to understand how deadenylation is regulated both globally and on speciﬁc mRNAs in response to environmental cues. mRNA decapping Nucleases of decapping and 59 to 39 degradation: mRNA decapping is carried out by a complex of the Dcp1 and Dcp2 proteins and is inﬂuenced by several other factors (Table 2). Dcp2 is the catalytic subunit and is a member of the Nudix family of pyrophosphatases (Van Dijk et al. 2002; Steiger et al. 2003). Dcp2 cleaves the cap structure to release m7GDP and a 59 monophosphate mRNA (She et al. 2008). Dcp1 is an EVH family protein (She et al. 2004) that inter- acts with Dcp2 to promote its catalytic activity (Deshmukh et al. 2008; She et al. 2008). The ﬁrst 300 amino acids of Dcp2 are sufﬁcient to promote decapping (Dunckley and Parker 1999) and fold into a two-domain structure wherein the N-terminal domain interacts with Dcp1 and the Nudix domain is present in the C-terminal domain (residues 100– 245) (She et al. 2006, 2008). Dcp2 has a conserved region between 245 and 286, which contains binding sites for the Edc3 protein and possibly other decapping activators (Harigaya et al. 2010). Yeast Dcp2 has an extended C- terminal region that is not required for general mRNA decapping. Since Dcp2 can shuttle into the nucleus (Grousl et al. 2009) and this region has sites that can enhance tran- scription (Gaudon et al. 1999), one possibility is that this region plays some role in controlling transcription (Shalem et al. 2011). Dcp2’s catalytic mechanism is a typical Nudix family re- action wherein Mg++ ions coordinated by a set of glutamic acid residues promote catalysis (Dunckley and Parker 1999; Steiger et al. 2003; She et al. 2006). Dcp2 catalysis is pro- moted by the closing of the bi-lobed Dcp2 structure to create a more active enzyme and more stable substrate binding (Deshmukh et al. 2008; She et al. 2008; Floor et al. 2010). Dcp1 is thought to enhance decapping by promoting the formation of this closed and more active structure (Deshmukh et al. 2008; She et al. 2008). The Dcp1/Dcp2 holoenzyme or Dcp2 alone prefer longer mRNA substrates in vitro, which is consistent with Dcp2 containing an ex- tended RNA-binding site and having a reaction mechanism that consists of an initial binding to the substrate followed by sliding to the cap structure (Steiger et al. 2003). However,