Chapter 17 Messenger RNA Half‐Life Measurements in Mammalian Cells
Introduction
Regulation of mRNA turnover in the cytoplasm is important for controlling the abundance of cellular transcripts and, in turn, the levels of protein expression (for reviews, see Parker and Song, 2004, Wilusz, 2001). mRNA stability can be regulated at different levels. Under a given physiologic condition, mRNAs display a wide range of stability. For example, c‐fos proto‐oncogene transcript is degraded rapidly in the cytoplasm with a half‐life of 10 to 15 min (Shyu, 1989, Treisman, 1985), whereas globin mRNA is rather stable and has a half‐life of several hours in the same cells (Shyu et al., 1989). Although each individual mRNA has its intrinsic stability under a given condition, stability of an individual mRNA may change in response to a variety of extracellular stimuli. Examples include the autoregulated degradation of tubulin mRNA in response to changes in tubulin concentration (Yen et al., 1988), the iron‐dependent destabilization of transferrin receptor mRNA (Casey, 1988, Muellner, 1989), the DNA synthesis–dependent destabilization of histone mRNA (Pandey and Marzluff, 1987), and the stabilization of lymphokine mRNAs by costimulatory molecules (Lindsten et al., 1989). Thus, modulation of mRNA stability provides a powerful means for controlling gene expression during the cell cycle, cell differentiation, the immune response, as well as many other physiologic transitions.
In mammalian cells, the first major step that triggers mRNA decay is deadenylation (i.e., removal of the 3′‐poly(A) tail). All major mRNA decay pathways recognized in mammalian cells, including mRNA decay directed by AU‐rich elements (AREs) in the 3′‐untranslated region (UTR) (Chen and Shyu, 1995), decay mediated by destabilizing elements in protein‐coding regions (Grosset et al., 2000), nonsense‐mediated mRNA decay (NMD) (Chen and Shyu, 2003), decay directed by microRNAs (miRNAs) (Wu et al., 2006), and decay of stable mRNAs such as β‐globin mRNA (Loflin, 1999a, Shyu, 1991), are initiated with deadenylation. Mammalian deadenylation exhibits biphasic kinetics. During the first phase, PAN2 poly(A) nuclease, presumably complexed with PAN3, shortens the poly(A) tails to ∼110 A nucleotides (nt) (Yamashita et al., 2005). In the second phase, the CCR4‐CAF1 poly(A) nuclease complex further shortens the poly(A) tail to oligo(A) (Yamashita et al., 2005). Decapping mediated by the DCP1–DCP2 complex was found to occur after either the first or the second phase of deadenylation (Yamashita et al., 2005). The RNA body can be degraded by the exoribonuclease XRN1 from the 5′‐end after decapping (Parker and Song, 2004). Alternately, the mRNA body can also be degraded from the 3′‐end after deadenylation by a large protein complex termed the exosome (Parker and Song, 2004).
To unravel the underlying processes of regulated mRNA turnover, a detailed analysis of the major components involved in mRNA turnover is required. The observation that deadenylation is the major trigger for cytoplasmic mRNA degradation in mammalian cells underscores the necessity of explaining the deadenylation step in the process of mRNA turnover. Thus, determination and characterization of the decay mechanisms demand that mRNA decay kinetics and precursor‐product relationships be accurately and readily monitored experimentally. The primary emphasis of this chapter is to describe two inducible promoter systems as examples to illustrate how mRNA turnover may be optimally investigated in mammalian tissue culture cells with transient transfection systems. Detailed step‐by‐step protocols are given so that the half‐lives of mRNAs of interest can be determined with experimental systems described here.
Section snippets
General Considerations of mRNA Half‐Life Measurements
Messenger RNA stability is often studied indirectly by monitoring changes in the steady‐state level of mRNA in the cytoplasm. However, changes in mRNA abundance are not necessarily caused by alterations in mRNA stability. For example, mRNA biogenesis in the nucleus (such as transcription, RNA processing, and/or mRNA export) may be fortuitously altered because of changes of the physiologic condition or in response to the environmental stimuli. Thus, alterations in the steady‐state level of mRNA
Determining mRNA Decay Constant
The turnover rate or stability of mRNA in vivo is usually reported as the time required for degrading 50% of the existing mRNA molecules (i.e., the half‐life of mRNAs). Before the half‐life of a given message can be calculated, the decay rate constant must be determined. Assuming an ideal in vivo situation, in which transcription of the mRNA of interest can be turned off completely (or at least to an undetectable level), mRNA decay follows first‐order kinetics. The rate of disappearance of mRNA
General inhibition of transcription
A relatively simple way of analyzing mRNA kinetics involves blocking cellular transcription with inhibitors that include actinomycin D (which interferes with transcription by intercalating into DNA) or 5,6‐dichloro‐1β‐1‐ribofuranosylbenzimidazole (DRB) (which interacts directly with the RNA polymerase II transcription apparatus) (Harrold et al., 1991). Typically, either actinomycin D (at a concentration of 5 to 10 μg/ml) or DRB (at a final concentration of 20 μg/ml) is added to cells, and the
Concluding Remarks
We have described two transcriptional pulsing methods that result in the synthesis of an mRNA population nearly homogeneous in size. These methods have several advantages over other approaches used to measure mRNA half‐life. They offer the opportunity to determine deadenylation and decay kinetics, as well as the precursor‐product relationship of mRNA turnover (Yamashita et al., 2005). Although the c‐fos promoter system is convenient to quickly address the mechanistic steps involved and
Acknowledgments
We thank many past and present members in our laboratory who have contributed in various ways over the years to the development of the approaches and protocols described in this chapter. The work was supported by National Institutes of Health (GM 46454) and in part by the Houston Endowment, Inc., and the Sandler Program for Asthma Research (to A.‐B. S.).
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