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BIOCHEMICAL CHARACTERIZATION OF THE ENDONUCLEASE PMR-1

DISSERTATION

Presented in Partial Fulfillment of the Requirements for

the Degree Doctor of Philosophy in Molecular, Cellular, and Developmental

Biology Program of The Ohio State University

By

Mark Nils Hanson B.S.

The Ohio State University 2001

Dissertation Committee: Approved by Professor Daniel R. Schoenberg, Adviser

Associate Professor Tien-Hsien Chang

Associate Professor Michael C. Ostrowski Advisor Assistant Professor Kathleen Boris-Lawrie Molecular and Cellular Biocherfiisty UMI Number 9999394

UMI

UMI Microform 9999394 Copyright 2001 by Bell & Howell Information and Learning Company. All rights reserved. This microform edition is protected against unauthorized copying under Title 17, United States Code.

Bell & Howell Information and Learning Company 300 North Zeeb Road P.O. Box 1346 Ann Arbor, Ml 48106-1346 ABSTRACT

The estrogen induced destabilization of serum protein mRNAs in Xenopus

liver occurs concurrently with the activation of the sequence selective, polysome-

associated endoribonuclease PMR-1. This endonuclease was cloned and found

to be a novel member of the peroxidase gene family. PMR-1 shares no

structural or sequence similarity with the known superfamily of vertebrate

ribonucleases, but does have a 57% sequence identity with human

myeloperoxidase (hMPO). PMR-1 was identified based on its ability to

preferentially cleave albumin mRNA within the single-stranded element APyUGA.

In this study I have: 1) demonstrated that unlike hMPO, PMR-1 does not have a

heme group, does not possess an autolytic activity when heated, is not a , and was shown to have a different enzymatic activity than hMPO.

2) shown that PMR-1 is widely expressed in many tissues and organisms. 3) determined that PMR-1 is a phosphoprotein, but phosphorylation of PMR-1 is not necessary for, and does not change with activation of the . 4) described the development of a sensitive ligation-mediated PGR amplification technique and its use in demonstrating the estrogen-induced appearance of albumin mRNA decay intermediates corresponding to in vivo cleavage by PMR-1.

11 Dedicated to ail of the people who supported me through this work.

I l l ACKNOWLEDGMENTS

I wish to express my appreciation and gratitude to my advisor, Dr. Daniel

R. Schoenberg, for his intellectual support, encouragement, patience, and enthusiasm. As my mentor he has had a great influence on my love of research and desire to leam. It is these intangibles that have been shaped the most by him.

I would also like to thank the members of my dissertation committee. Dr.

Kathleen Boris-Lawrie, Dr. Tien-Hien Chang, and Dr. Michael Ostrowski for their support and stimulating discussions.

I want to thank the Ohio State University and the Molecular, Cellular, and

Developmental Biology Program for its support.

I would like to thank all of my colleagues who have educated and inspired me, including Lynn, Jenni, Xiaoping, Haidong, Ping, Joy, Kirsten, Venk, Rob,

Jing, Changhong, Feng, Katie, Sanjay, Usha, Julie, Dr. Jones, and most of all

Elena for the patience two teach me so much, and Kris for always being there.

I would like to thank my father Dr. John Nils Hanson, mother Stephanie

Morgan Hanson, sister Laurel Hanson, and Susan Brkich for all the love and support that made the rough spots smoother.

IV VITA

March 17, 1972 ...... Bom Mt. Lebanon, PA, USA

1990-199 4...... B.S. Gettysburg College

1991-presen t ...... Graduate Teaching and Research Associate, The Ohio State University

PUBLICATIONS

1) Hanson, M.N., and Schoenberg, D.R., “Identification of in vivo mRNA decay intermediates corresponding to sites of in vitro cleavage by polysomal ribonuclease 1." Journal of Biological Chemistry, In Press. (2001)

2) Cunningham, K.S., Hanson, M.N., Schoenberg, D.R., “Polysomal Ribonuclease 1 exists in a latent form on polysomes prior to the estrogen activation of mRNA decay.” Nucleic Acid Research, In Press (2001)

3) Cunningham, K.S., Hanson, M.N., Schoenberg, D.R. ‘The mRNA endonuclease PMR-1” Methods in Enzymology. In Press. (2001) N.B.: Co-first author with Cunningham.

4) Chemokalskaya, E., Dubell, A.N., Cunningham, K.S., Hanson, M.N., Dompenciel, R.E., Schoenberg, D.R., “A polysomal ribonuclease involved in the destabilization of albumin mRNA is a novel member of the peroxidase gene family.” RNA 4:1537-1548. (1998) N.B.: Co-first author with Chemokalskaya, Dubell and Cunningham.

FIELDS OF STUDY

Major Field: Molecular, Cellular, and Developmental Biology TABLE OF CONTENTS

Page

ABSTRACT...... ii

DEDICATION...... iii

ACKNOWLEDGMEMTS...... iv

VITA...... V

TABLE OF CONTENTS...... vi

LIST OF TABLES...... x

LIST OF FIGURES...... xi

ABBREVIATIONS...... xlll

CHAPTERS:

1. INTRODUCTION...... 1

1.1 The role of mRNA half-life in gene expression ...... 1 1.2 Cis and Trans action stability determinants...... 3 1.3 mRNA decay in prokaryotic cells ...... 8 1.3.1 Endonucleases ...... 8 1.3.2 Exonucleases ...... 10 1.3.3 Degradosome ...... 12 1.4 mRNA decay in Saccharomyces cervisiae...... 14 1.4.1 5’-3’ decay: deadenylation-decapping ...... 14 1.4.2 3’-5’ decay: the exosome ...... 19 1.5 mRNA decay in mammalian cells ...... 22 1.5.1 The role of AU-rich elements (ARE) in mRNA decay... 22 1.5.2 3’-5’ exonucleolytic decay in mammalian cells ...... 28 1.5.3 mRNA surveillance in mammalian cells ...... 30 1.6 mRNA endonucleases in vertebrate cells ...... 34 1.7 Estrogen regulation of mRNA stability ...... 37 1.8 Identification and characterization of PMR-1 ...... 38

vi 1.9 Human myeloperoxidase ...... 41 1.10 Purpose of this study ...... 42 1.10.1 Biochemical comparison of PMR-1 and hMPO ...... 42 1.10.2 Expression of PMR-1 in Xenopus and murine tissues and Estrogen phosphorylation of PMR-1 ...... 43 1.10.3 In vivo mapping of endonuclease cleavage sites ...... 44

2. MATERIALS AND METHODS...... 48

2.1 Purification of PMR-1 from Xenopus liver ...... 48 2.2 Crude liver polysome isolation ...... 48 2.3 FPLC purification of PMR-1 ...... 49 2.4 Bradford assays ...... 51 2.5 Autolytic cleavage of hMPO ...... 51 2.6 Western blotting ...... 51 2.7 Spectrum of hMPO and PMR-1 ...... 52 2.8 Endoglycosidase H cleavage ...... 53 2.9 Concanavalin A binding ...... 53 2.10 HMPO ribonuclease activity ...... 54 2.11 Peroxidase activity assay ...... 54 2.12 In vitro activity assays ...... 55 2.13 Purification of Liver RNArGuanidinium-lsothiocynate method ...... 55 2.14 Purification of total RNA from cell culture using TRIzol® ...... 57 2.15 Isolation of total protein from frog tissues ...... 57 2.16 Isolation and maturation of Xenopus Oocytes ...... 58 2.17 Isolation of total protein from mouse tissues ...... 59 2.18 Activity assays with frog and mouse tissues ...... 59 2.19 Calf intestine alkaline phosphatase treatment of protein samples ...... 60 2.20 Isolation of total protein from cell culture ...... 60 2.21 Ligation Mediated Polymerase Chain Reaction (LM- PCR)...... 61 2.21.1 Primer ligation: DMSO method ...... 61 2.21.2 Primer ligation: PEG method ...... 61 2.21.3 RT-PCR...... 62 2.21.4 Product recovery, reamplificaiton and sequencing ...... 63 2.22 In vitro activity assay ...... 65 2.23 RNase T1 digestion ...... 65 2.24 Isolation of cytoplasmic protein from hepatocytes ...... 66 2.25 Xenopus primary hepatocyte cell culture ...... 66 2.26 Transfection of hepatocytes with antisense Morpholino’s ...... 68

V ll 2.27 Isolation of Xenopus hepatocyte cytoplasmic/nuclear RNA...... 69 2.27.1 Northern blot ...... 69 2.27.2 Hybridization of Northern blot ...... 70 2.27.3 Labeling probe ...... 71

3. RESULTS...... 73

3.1 Biochemical comparison of hMPO and PMR-1 ...... 73 3.1.1 The absence of heme distinguishes PMR-1 from hMPO...... 73 3.1.2 hMPO is glycosylated whereas PMR-1 is not ...... 81 3.1.3 PMR-1 and hMPO have distinct enzymatic activities... 89 3.2 Expression of PMR-1 in other tissues and organisms.. 92 3.2.1 PMR-1 antibody cross reacts with a protein in Xenopus and murine tissues ...... 92 3.2.2 Unlike murine tissues, Xenopus tissues all produced the major doublet cleavage of PMR-1 ...... 96 3.3 Estrogen and development do not change the effect of CIAP treatment on PMR-1 ...... 101 3.4 Development of LM-PCR for mapping of in vivo degradation intermediates ...... 107 3.4.1 A sensitive LM-PCR assay for detecting in vivo mRNA degradation intermediates ...... 107 3.4.2 Identification of in vivo albumin mRNA decay intermediates corresponding to sites cleaved in vitro by PMR-1...... I l l 3.4.3 Application of LM-PCR to identify degradation intermediates from the 3’ end of albumin mRNA ...... 118 3.4.4 Impact of secondary structure on PMR-1 cleavage of the albumin mRNA 3’ end ...... 124 3.4.5 Identification of PMR-1 cleavages within the vigilin binding domain of vitellogenin mRNA ...... 127 3.4.6 Identification of decay intermediates consistent with in vivo endonuclease cleavage within the c-myc CRD...... 130 3.5 PMR-1 antisense Morpholino has no effect on levels of albumin mRNA in primary hepatocytes ...... 133

4. DISCUSSION...... 141

4.1 PMR-1 and hMPO are biochemically different from each other ...... 141

VIll 4.2 PMR-1 is expressed in multiple Xenopus and murine tissues ...... 146 4.3 The effects of CIAP treatment does not change with estrogen or development ...... 149 4.4 Development of PM-PGR method ...... 150 4.4.1 LM-PCR identifies the same cleavages in vivo as . previously mapped in vitro...... 151 4.4.2 In vivo mapping of the 3’ portion of albumin is consistent with PMR-1 as the major factor involved in the degradation of albumin mRNA ...... 153 4.4.3 LM-PCR suggests that PMR-1 is also involved in the degradation of vitellogenin mRNA ...... 154 4.4.4 LM-PCR can also be applied to rare mRNAs such as c-myc...... 155 4.5 Morpholino antisense PMR-1 oligos have no effect on the estrogen destabilization of albumin in hepatocytes ...... 156 4.6 Concluding remarks ...... 157 LIST OF REFERENCES...... 161

IX LIST OF TABLES

Table Page

1.1 Cis and Trans stability determinants ...... 6

1.2 Major trans factors involved in E. coli mRNA degradation ...... 9

1.3 Major trans factors involved in S. Cerevisiae mRNA degradation 15

1.4 Major trans factors involved in vertebrate mRNA degradation ...... 23

3.1 Transfection efficiency of Morpholino oligo into primary hepatocytes ...... 137 LIST OF FIGURES

Figure Page

1.1 Identification of in vivo cleavage products by primer extension ...... 45

3.1 Sequence and structural features of PMR-1 ...... 74

3.2 PMR-1 does not undergo autolytic cleavage upon heating ...... 77

3.3 Wavelength scans demonstrate that unlike hMPO, PMR-1 does not have a heme group ...... 79

3.4 N-linked glycosylation sites on hMPO and PMR-1 ...... 83

3.5 Endoglycosidase H digestion shows no high on PMR-1 ...... 85

3.6 Concanavalin A-Sepharose binding shows no oligosaccharides on PMR-1 ...... 87

3.7 Comparison of the enzymatic activities of hMPO and PMR-1 ...... 90

3.8 Western blot of frog and mouse tissue shows PMR-1 is present in many tissues ...... 94

3.9 Activity assay of frog tissue proteins demonstrates the presence of active PMR-1 in all tissues ...... 97

3.10 Activity assay of murine tissue proteins suggests that PMR-1 is expressed in mammals ...... 99

3.11 CIAP treatment of PMR-1 alters its mobility on SDS-PAGE ...... 103

3.12 Development has no effect on the ability of CIAP treatment to alter the mobility of PMR-1 on SDS-PAGE ...... 105

3.13 The LM-PCR protocol for identification of mRNA decay intermediates ...... 109

XI 3.14 Optimization of the ligation protocol ...... 112 3.15 LM-PCR identification of decay intermediates in the 5’ coding region of albumin mRNA ...... 115

3.16 LM-PCR identification of decay intermediates in the 3’ end of albumin mRNA ...... 119

3.17 In vitro cleavage of the 3’ end of albumin mRNA ...... 122

3.18 Secondary structure of the 3’ end of albumin mRNA ...... 125

3.19 LM-PCR identification of decay intermediates in the vitellogenin 3’ UTR following estrogen withdrawal ...... 128

3.20 LM-PCR identification of degradation intermediates within the c-myc coding region determinant ...... 131

3.21 Transfection efficiency of Morpholino oligo into primary hepatocytes ...... 135

3.22 Northem blot of total hepatocyte RNA from Morpholino treated cells ...... 138

4.1 Predicted tertiary structure of PMR-1 ...... 144

XU ABREVIATIONS

a-PMR-1 - anti-PMR antibody ABP - ARE binding protein AoNPV - Autographe oaUfomica nuclear poly bed rosis virus Apo II — apolipoprotein II AB 1277/Ab 2/ Ab 4 - antibodies 1277/2/4 ARE - AU-rich element ATP - adenosine triphosphate AU-rich — adenosine-uracil rich AYUGA — A-pyrimidine-UGA B1UTR-15 - vitellogenin 81 3’UTR p-AR - p-adrenergic receptor P-ME - p-mercaptoethanol bp — base pair c a t- choramphenicol acetyl cDNA — complementary DNA CLB — crosslinking buffer CRD — coding region determinant CRD-BP — coding region determinant binding protein CRE - cytosine-rich element OTP - c^osine triphosphate dCTP - deoxycytosine triphosphate DAN - deadenylating nuclease DEPC - diethyl pyrocarbonate Dl - defective interfering particle DMSO — dimethyl sulfoxide DNase — deoxyribonuclease DNA - deoxyribonucleic acid DSE - downstream element DTT - dithiothreitol elF4E/G/F - eukaryotic initiation factors 4 E/G/F E. coli — Escherichia coli EDTA - ethylenediaminetetraacetic acid EGTA - ethylene glycol-bis(B-aminoethyl Ether) N,N,N’,N’-tetraacetic acid ELAV - embryonic lethal abnormal visual ER - estrogen receptor

x i i i eRF1/eRF3 - eukaryotic release factor 1/3 ErEN — Erythroid-enriched endonuclease FBS - fetal bovine serum G3BP — RasGAP Src homology 3 binding protein GC-rlch - guanoslne-cytoslne rich GTP — guanoslne triphosphate 3 -OH - 3’-hydroxyl group hMPO - human myeloperoxidase IgG - Immunoglobulin IL-2/IL-3/IL-6 - Interleukin 2/3/6 IRE — Iron response element I RE-BP - Iron response element binding protein Kd — dissociation constant KH domain - K homology domain ^"’G-cap - 7-methyl guanoslne cap ^"’GDP — 7-methyl guanoslne diphosphate ^"’GMP - 7-methyl guanoslne monophosphate GM-CSF — granulocyte-macrophage colony stimulating factor hsp70 - heat shock protein 70 Lsmp - Sm-llke protein LSM-I - Lsm complex MAP kinase — mitogen activated kinase MBU — molecular biology unit MIE - MATal Instability element MOI — multiplicity of Infection mRNA - messenger ribonucleic acid mRNase - messenger ribonuclease mRNP — messenger ribonucleoprotein particle NMD - nonsense mediated decay NP-40 - Nonldet P-40 (detergent) nt - nucleotide ollgo-dT - thymine oligonucleotide PABP - poly(A) binding protein (vertebrates) Pabp - poly(A) binding protein (yeast) PAN - poly(A) nuclease PAP1 -poly(A) polymerase 1 (bacteria) PARN - poly (A) ribonuclease PBS - phosphate buffered saline PCR - polymerase chain reaction PEI GO - polysomal salt extract PGK1 - phosphoglycerate kinase 1 pi - Isoelectric point PMR-1 - polysomal ribonuclease 1 PMSF - Phenylmethylsulfonyl fluoride PNPase - polynucleotide phosphorylase

x i v POL III — RNA polymerase III PVDF — Polyvinylidene fluoride RasGAP - GTPase activating protein (for ras) RER — rough RLI - RNase L Inhibitor rRNA - ribosomal RNA RNA — ribonucleic acid RNase(s) — rlbonuclease(s) RNP(s) - ribonucleoprotein RPM — revolutions per minute RRM - RNA recognition motif SI 00 - 100, 000 X g supernatant 8130 - 130, 000 X g supernatant S. cerevisiae — Saccharomyces cerevisiae SDS — sodium dodecylsulfate SDS-PAGE — sodium dodecylsulfate polyacrylamide gel electrophoresis Sf9 — Spodoptera frugiperda 9 SKI - super killer snRNA - small nuclear RNA snoRNA(s) — small nucleolar ribonucleic acld(s) SSG — sodium chlorlde/sodlum citrate SV40 - simian virus 40 TBS - Tris-buffered-sallne TBST - Tris-buffered-sallne Tween-20 TfR — transferrin receptor TNF-a - tumor necrosis factor a tRNA — transfer RNA uORF - upstream open reading frame DTP - uracil triphosphate UV - ultraviolet 5’UTR — 5’ untranslated region 3’UTR - 3' untranslated region v/v — volume per unit volume vhs - viral host shutoff vit. - vitellogenin w/v — weight per unit volume W P100 — washed polysomes X al 60 — Xenopus albumin RNA 160 Xa470 - Xenopus albumin RNA 470 XLN1 - Xenopus laevis nuclease 1

Units |ig/ml - microgram per millilitre pg - microgram pM - micromolar

XV p.Ci — microcurie |ii — microliter jiJ/cm^ - microjoules per square centimeter cm - centimeter fmol - femtomole g — gram g/L — grams per liter g/ml — grams per millilitre kb - kilobase kDa — kilodalton mA - milliampere ml — milliliter ml/g — milliliters per gram mg — milligram M — molar mM — milimolar mmol — milimole nm - nanometer pmol - picomole V — volt V/cm - volts per centimeter

XVI CHAPTER 1

INTRODUCTION

1.1 The Role of mRNA half-life in gene expression.

The end result of gene expression Is the production of a protein. The accumulation of this protein product is regulated at many different steps, including transcription, messenger RNA (mRNA) half-life, , and protein half-life. Regulating the amount of protein can affect how a cell grows, differentiates, and responds to its environment (1). This dissertation will focus on an enzyme that affects the half-lives of several mRNAs.

Change in mRNA concentration at any time is a pseudo first-order process, and is dependent on the amount of mRNA that is present (1 ). The half- life of a mRNA can affect both the rate of disappearance and the rate at which a new steady state level can be achieved. Yeast mRNA half-lives typically range between 1 and 30 minutes while mammalian mRNA half-lives can range from 5 minutes to hundreds of hours (2). A relatively small change in the half-life of an mRNA (2-4 fold) can drastically reduce the amount of a message present. For instance, compare a half-life of 10 minutes versus 40 minutes, a mere 4 -fold difference. If you start with 10,000 molecules per cell, after 2 hours the mRNA

1 with a 10 minute half-life will have virtually disappeared. The mRNA with a 40 minute half-life will still be relatively abundant with over 1,000 molecules left. The other major effect is the rate of change following a change in transcription.

Assume that mRNA’s A and B have the same change in the rate of transcription but mRNA A is 10 fold less stable, it will reach its new steady state 10 times more quickly.

The stability of a mRNA can be regulated through sequence and structural cis elements that interact with trans acting factors such as RNA binding proteins and endoribonucleases. It is the interaction of these cis and trans elements that impart a half-life on a message that is unique to the cell type, and the environmental stimuli present. This allows for a message to have very different half-lives in different cell types under different conditions. These stability determinants range from common features such as the poly(A) tail and 5’ cap to specific sequences that may only be present on a transcript or class of transcripts such as the iron response element (IRE). Some messages may not have any specific degradation signals while others like c-myc will have several that interact to tightly regulate its expression. Once degradation is commenced it appears to be completed rapidly. This suggests that the interplay between these cis and trans acting stability elements is an important factor in the process of mRNA decay (3). Although mRNA half-life plays an important role in determining the

expression of its gene product, Gygi et. al. (4) have demonstrated in yeast that there is no strong correlation between the amount of mRNA and the amount of

protein present in a cell, it was shown that the amount of protein expressed

could vary by 20-fold with the same amount of mRNA being present. The

amount of mRNA could vary 30-fold with the same amount of protein being

expressed. Although these numbers are probably exaggerated due to the limits

of the technique used, they do demonstrate the point that one cannot directly

predict the amount of protein by looking at the amount of mRNA and vice versa.

It is important to remember that even though mRNA levels do not directly mirror

protein levels mRNA degradation still plays an important role in the regulation of protein expression as discussed above.

1.2 Cis and Trans acting stability determinants.

Most often cis acting stability determinants are identified by a “cut and paste” method where hybrid transcripts are created that have a reporter such as cat or /uc fused to portions of the mRNA under study. The stability of the hybrid transcript is then measured. Once a region is identified as a potential stability determinant, deletion analysis and/or linker scanner analysis is performed to narrow down the element and demonstrate that it is sufficient to confer the wild type effect on mRNA stability. However, it is important to be aware of the possibility of disrupting multiple stability determinants that are necessary to impart the expected half-life. Also, there are many variables that can indirectly

3 effect the stability of a transcript. These variables include primary and secondary structure, translational efficiency, and intracellular localization. For example one might find that a particular deletion stabilizes a transcript not because of the loss of a destabilizing element but by an increase in translation efficiency (1 ).

Cis acting stability determinants can often be position dependent. For example both the Mata1 instability element (MIE) in yeast and the c-myc coding

region determinant (CRD) in mammalian cells have several rare codons, and the insertion of a stop codon upstream of either element leads to a stabilization of the transcript (5). This suggests that translation through the element is required for destabilization of the transcript. Also in yeast the presence of an upstream

reading frame uORF may also destabilize a transcript, although it does not always have this effect. Both the YAP2 and PPR1 uORFs have been shown to naturally destabilize there transcripts. Fusion of the PPR1 uORF upstream of the very stable PGK1 coding region makes it a very unstable transcript (3). Also, a single nucleotide mutation that introduces a seven codon uORF in the caf mRNA destabilizes the message four-fold. Table 1.1 lists the major cis and trans acting factors involved in mRNA stability that have been identified to date.

Trans acting factors can effect a single transcript, a class of transcripts, or be involved in general mRNA decay. These factors can"be further subdivided into those that bind directly to the mRNA such as poly(A) binding protein (PABP), eukaryotic initiation factor 4E (elF4E), and AU-rich element (ARE) binding

4 proteins, and those factors that do not bind directly to the mRNA such as tubulin,

histone, and heat shock proteins (1). Trans acting factors can act as either a

stabilizing or destabilizing factor for the mRNA they interact with. Some factors

such as AUF1 can function as both depending on which protein complex the

factor is associated with. AUF1 interaction with either the a-globin mRNA, a

stabilizing effect, or c-myc mRNA, a destabilizing effect, is an example of these

duel functions (6,7).

Trans acting factors can either act alone or as part of a larger complex.

For example, the a-complex which consists of the poly(C) binding proteins aCP1

and aCP2, as well as AUF1 binds to the 3-UTR of a-globin mRNA stabilizing the

transcript preventing cleavage by the endonuclease ErEN (6,8). The poly(C)

binding proteins have been demonstrated to interact with PABP in a way that

mutually stabilizes binding of the a-complex to the cytosine rich element (CRE)

and PABP to the poly(A) tail. Deadenylation, which leads to loss of PABP

binding, has been shown to disrupt binding of the a-complex and therefore lead to destabilization of the transcript (9).

Translation has been demonstrated to be an integral part of mRNA stability. First, a number of nucleases are associated with the translation machinery. Second, translation can remove protective proteins from the transcript, as well as alter the secondary structure of the mRNA potentially rendering instability determinants susceptible to binding by

5 cis trans mRNA elements elementsRef Comments

P21 ? HuR (10) UV light regulated/3'UTR P21 ? ? (10) tyrosine phosphorylation mediated stability/UV ind. P21 ARE 24 and (11) alphal -agonist phenylephrine activates 52 kDa binding GM-CSF ARE hnRNP (12) phosphorylation/human T-lymphocytes A1 c-fos/GM- ARE? (13) req. association with polys for degradation CSF TNF- ARE 55 kDa (14) LPS mediated/3'UTR alpha TNF- ARE T ristetrap (15) GOGH family of zinc finger proteins alpha rolin (TTP) ? ARE ELAV-like (16) binds ARE + poly(A) tail/3'UTR papiloma ARE HuR (17) 3’UTR/hnRNP G1 and G2 also bind 1 late GAPDH ARE GAPDH (18) req. NAD+ binding domain/found on polys IGFBP-1 ARE ? (19) 3’UTR/change in stability with developmental stage Beta-AR AU-rich BARB=H (20) both HuR and hnRNP A1 bind 3’UTR uR mucin AU-rich ? (21) ARE destabilized in lower eukaryote family (trypanosome) bcl-2 AU-rich ? (22) apoptosis includes decay apoB AU-rich apobec-1 (23) not ARE/apobec=cysteine diaminase of editing enz. 7alpha- AU-rich ? (24) ARE’s not responsible for bile acid regulation hydroxyla se hFGF-2 AU-rich ? (25) non ARE/2 X 122 nt repeat/alternate poly(A) IL-2 AU- 55 kDa (26) protects from endo cleavage in 3’UTFVJurkat rich/160 nt cells IL-2 5’UTR/3’U JNK (27) JNK activity through 55’UTR yet 3’UTR also TR mediated necessary PAI-1 A-rich/U- 38(U)/5G, (28) muliti-protein complex forms on 134 nt A-rich rich 61.76(A) region PAI-1 3’UTR ? (29) cAMP med./3’ most 134nt of 3’UTR are (134nt) sufficient

Table 1.1 : Cis and Trans stability determinants continued continued

apoll 3’UTR 34 and (30) two protein binding sites that regulate 60 kDa stability apoll 3’UTR endonucl (31) endo cleavage in ssRNA/requires context of (AAU/UAA) ease mRNP cyciInA/R 3’UTR p82 (32) p82 phos. by S/T kinase necessary for R unmasking LDH-A 3’UTR/U- 96/67/52/ (33) PKA/cAMP-medlated/critical stem-loop rich 50 kDa LDH-A 22 nt ? (34) identification of cAMP-stablliziong region (CSR) (GSR) APP 29 nt hnRNPC/ (35) 3’UTR/reticulocyte lysate nucleolln APP 5’ÜTR (90 ? (36) IL-1 stabilization/translational enhancer nt) AT-R 5’UTR ? (37) cAMP/PKA-mediated down regulation/non- 3’UTR AC914 5’UTR 40 S (38) Dictyostelium/PKA dep. Activity on S6 reg stability. lL-11 5’ÜTR-CR- ? (39) PKG mediated stability multiple cis regions 3’UTR necessary uPAR 51 nt uPARbp (40) MS-1 cells/PMA/coding region vitell/dystr 75 nt vlgilin/15 (41) estrogen stabilized/3’UTR iphin 5 kDa VEGF 126 bp (21 hnRNP (42) 3’UTR/hypoxia bp) L/60 kDa Collagen stem-loop 120 kDa (43) near AUG/alphaGP in 3’UTR binds 120 kDa alpha 1 GLUT1 105 nt 37 and (44) 3’UTR/TNF-a regulated/protects from endo 40 kDa activity GLUT4 200 nt 40 S/? (45) preinitiation complex forms+ inc. binding to 3’UTR SAUR- ? ? (46) 3’UTR/in plants AC1 GM-GSF SLDE ? (47) not ARE/stem-loop destabilizing element/3’UTR COX-2 ? ? (48) p38 MAPK dependent stabilization c-myc 3’UTR RasGAP/ (49) G3BP has nuclease activity dependent on G3BP phos. c-myc ORD GRD-BP (50) GRD-BP binding protects from endo cleavage

Table 1.1 : Cis and Trans stability determinants nucleases or binding proteins. Third, translation could localize the mRNA to a subcellular region that has the degradation machinery. Finally, translation itself may be an active part of the destabilization of a mRNA (51). The impact of translation on mRNA decay will depend on the particular system in question and the metabolic state of the cell.

1.3 mRNA decay in prokaryotic cells.

There have been over 20 RNases discovered in Escherichia coii {E. coii) but only 6 of these are currently known to function in mRNA decay. The rest of the RNases, such as RNase P, D, PH, T, and BN have a role in the processing and degradation of tRNA and rRNA (52,53). Even though prokaryotes have relatively unstable mRNA compared to eukaryotes, the half-life of a RNA can still be changed significantly. Table 1.2 lists the known factors involved in decay of prokaryote mRNA decay.

1.3.1 Endonucleases.

The major endonuclease involved in E. co//mRNA degradation is RNase

E, which is also involved in the processing and degradation of rRNA. This enzyme is quite large at 118 KDa and has been shown to have multiple functional domains. Only about 60% of the N-terminus is needed for the endonuclease activity while the C-terminal portion of the enzyme is involved with protein interactions which will be addressed more in section 1.3.3. RNase E has a loose specificity in that it cleaves 5’ to an AU dinucleotide that is in a single

8 Factor Known Function

RNase III endonuclease cleaves dsRNA RNase G (CatA) endonuclease cleaves ssRNA RNase II 3’-5’ exonuclease oligoribonuclease decay 2-5 oligonucleotides to mononucleotides

Deoradosome (54)

RNase E Endonuclease cleaves 5’ to ssAU dinucleotide PNPase polynucleotide phosphorylase RhIB RNA helicase enolase energy generation? GroEL assemble or folding? DnaK polyphosphate kinase formation of polyphosphate chains

Table 1.2 Major trans factors Involved InE. co// mRNA degradation. stranded conformation. Interestingly, purified RNase E can also shorten 3’

poly(A) and poly(U) tails with an exonuclease like activity that requires the same

N-terminal portion of the enzyme as the endonuclease activity (54). Knockout

mutation of the gene that encodes RNase E, me, is lethal and several temperature sensitive mutants have been identified (55).

The other prominent endonuclease is RNase III which cleaves in double

stranded regions of some mRNA’s. Although RNase III does initiate the

degradation of these messages, it’s knockout has a mild phenotype suggesting that it does not have a major role in mRNA decay (56).

The third endonuclease identified to play a role in mRNA decay is RNase

G or CafA. This enzyme has extensive sequence similarity to the N-terminal half

of RNase E (55), and also has a specificity that overlaps with but is distinct from

RNase E. Interestingly, non-functional mutations of this gene are not lethal, but do decrease the viable temperature of a temperature sensitive me-1 mutant (57).

1.3.2 Exonucleases.

All three exonucleases discovered to function in mRNA degradation in E. co//are 3’-5’ exonucleases. Which is intriguing since the net degradation occurs in a 5’-3’ direction. This is believed to be due to the fact that RNase E prefers to have a free 5’ end for its activity, therefore cleaving messages toward the 5’ end first and moving consecutively toward the 3' end. The exonucleases then

1 0 degrade these cleavage products in a 3’-5’ fashion producing a net 5-3’ degradation of the mRNA (58,54).

PNPase degrades single stranded RNA from the 3’ end by reversible phosphorolysis creating nucleoside 5’ diphosphates. RNase II, on the other hand, is a hydrolytic enzyme that removes nucleoside 5’ monophosphates from the 3’ end of a mRNA. RNase II accounts for about 90% of the exonuclease activity in the gram negative bacteria E. coii. However, it is just a minor part of the activity in gram positive bacteria such as Bacillus subtilis. Despite it’s prominent role in mRNA decay, deletion of the gene encodingRNase II is not lethal, but co-deletion with PNPase is (59). Both exonucleases are sensitive to secondary structure which is surprising considering how prevalent strong secondary structure is in phage, plasmid, and bacterial mRNA. Rho-independent termination (60), REP (repetitive extragenic palindrome) sequences (61), and processing by RNase III all leave quite stable stem loops at the 3’ end of the

RNA. This issue will be addressed more in depth in section 1.3.3.

Oligoribonuclease which is found in a homodimer complex in E. call decays small 2-5 oligonucleotides to mononucleotides. This activity is essential for cell viability as demonstrated by knockout mutations (62). When the am gene that encodes for oligoribonuclease is deleted an accumulation of oligonucleatides between 2-5 residues long is seen. Genes with close homology to am have been found in organisms all the way up to humans.

11 1.3.3 Degradosome.

All of the involved in mRNA degradation in E. coH, with the exception of RNase II, RNase III, and oligoribonuclease are present in a large macromolecule complex called the degradosome, which is between 1.5-2.4 x 10® daltons. The core of the degradosome is RNase E, which has a C-terminal half that includes binding sites for RhIB (DEAD box ATP-dependant helicase), enoloase (glycolytic enzyme), PNPase, and RNase E. Although the exact composition of the degradosome is not known, predictions are that it is composed of at least one dimer of RNase E, one trimer of PNPase, two dimers of enolase, and one dimer of RhIB. This has been supported by Colbem et. a i in vitro where they were able to reconstitute the ATP-dependent degradation of

REP containing mRNA with a minimalistic degradosome (63). The presence of an endonuclease, exonuclease, and helicase in the same complex suggests that the degradosome may be important for the coordination of endo- and exonuclease digestion and the unwinding of RNA structures. Consistent with this, the degradosome is required for the degradation of highly structured mRNA

(63). Enolase, on the other hand, does not have a known function in the degradosome.

Other proteins that have been found to associate with the degradosome under different conditions are the heat-shock chaperone’s GroEL and DnaK, but no role in RNA degradation is known for these proteins. However, GroEL has

1 2 been found to associate with a temperature-sensitive but not wild-type RNase E.

This suggests a possible role for the chaperones in the assemble or folding of the degradosome (64). Polyphosphate kinase, an enzyme th at polymerizes the terminal phosphate of ATP into long polyphosphate chain s, is associated with the degradosome (65). Regulation of polyphosphate levels m ay control degradosome activity. Also, fragments of the 16S and 23S rRNA have been found associated with the degradosome. These are belietved to be products of

RNase E processing or degradation and not play a role in degradosome activity.

Although poly(A) polymerase I (PAP I) can associante with RNase E it has not been identified as associating with the degradosome, and the specificity of

RNase E on certain substrates is altered in a PAP I mutant strain (66). Thus, it would not be surprising if more proteins are found to interact with the degradosome as it degrades various RNA substrates.

A role for PAP I in RNA stability is seen in a poly(A])-deficient strain where the mRNAs in general are stabilized (67). The converse has also been seen where mRNA half-lives generally decrease with increased cellular poly(A) levels.

However certain transcripts which include the mRNAs that encode for the enzymes RNase E and PNPase are stabilized with increasing PAP I levels (68).

This stabilization of enzymes integral to general RNA degradation raised the possibility that regulatory controls exist that balance the m RNA decay and polyadenylation pathways. The major function of PAP I is to help the

13 degradation machinery, in particular PNPase, overcome strong secondary structures by adding a poly(A) tail that can be used to get the degradosome started and allow the helicase to unwind these structures (69).

1.4 mRNA decay in Saccharomyces cerevisiae.

The fission (budding) yeast Saccharomyces cerevisiae (S. cerevisiae) is the most abundantly studied eukaryotic organism for mRNA stability. In contrast to prokaryotes, eukaryotic mRNA requires extensive post transcriptional modifications such as splicing, capping, and polyadenylation. Interestingly all three of these processes can lead to more stable transcripts as discussed below.

The most important modifications are the cap and poly (A) tail both of which function to stabilize the mRNA since the degradation machinery has to pass through these structures to degrade the mRNA. Also, these structures can increase the efficiency of translational initiation via 5’end - 3’end communication

(70). Table 1.3 lists the known factors that are involved in the degradation of

RNA in Yeast.

1.4.1 5 -3' decay: deadenylation-decapping.

Work primarily done in Roy Parker’s lab has elucidated the major pathway of mRNA turnover in S. cerevisiae. This pathway has three major steps. The first is shortening of the poly(A) tail to oligo(A) lengths. The second is removal of the cap. The third is highly processive S’-3’ exonuclease degradation of the mRNA (73).

14 Factor (71 ) Known function

X m lp 5’-3’ exonuclease Xm2p (Hkelp) 5-3' exonuclease Panpl 3’-5’ exonuclease Dcpip Decapping Dcp2p Decapping Mtr4p RNA helicase PAN Poiy(A) nuclease (3’-5’ exonuclease) Ski2p RNA helicase Ski3p/Ski8p M rtip Upflp mRNA surveillance/ (ATPase/helicas Upf2p mRNA surveillance UpfSp mRNA surveillance

Exosome (72) Rrp4p 3’-5’ exonuclease Rrp40p 3’-5’ exonuclease Rrp41 p (Ski6p) 3'-5’ exonuclease phosphorylase Rrp42p 3’-5’ exonuclease Rrp43p 3’-5’ exonuclease Rrp44p (Dls3p) 3'-5’ exonuclease hydrolase Rrp45p 3’-5’ exonuclease Rrp46p 3’-5’ exonuclease Mtr3p Csl4p (Ski4p) RNA binding (particularly to mRNA) Rrp6p nuclear 3'-5’ exonuclease hydrolase Ski7p GTPase

Table 1.3 Major trans factors involved in S.cerevisiae mRNA degradation.

15 Surprisingly, mRNA deadenylases remain poorly characterized. The yeast genome encodes several potential deadenylases. However, pentuple deletion of these show no clear defect in deadenylation (71). In strains deleted of the PABP gene which encodes poly(A) binding protein, (Pabp) deadenylation is slowed, suggesting that Pabp stimulates deadenylation (74).

Contrary to deadenylation, the mechanism of decapping is relatively well known. Decapping produces ^"^GDP and a mRNA with a free 5’ monophosphate

(75). This can be accomplished in vitro by Dcpi p alone. However in vivo other proteins are needed for decapping such as Dcp2p, which converts ^"’GDP to

^"™GMP and phosphate. This is believed to prevent product inhibition (76).

Interestingly, Dcpip expressed in a Dcp2p mutant strain and purified is still inactive. While, Dcpip produce in a DCP2 cell, and separated from Dcp2p by a high salt wash was fully functional, suggesting that Dcp2p is needed for the production of active Dcpip but does not need to be present for Dcpi p to be active (76)(T. Dunckley, M. Tucker, and R. Parker, submitted for publication).

Parker’s lab has demonstrated that regulation of decapping occurs by a direct competition for cap binding between elF4E and Dcpip (77).

Another protein that seems to be required for in vivo activity is

Sbp8p/Lsm1 p which interacts with a complex consisting of the related proteins

Lsm2p-8p called the LSM1. This complex binds to the 3’ end of U6 small nuclear

RNA (snRNA) as a doughnut shaped ring (78). Lsm lp itself does not associate

1 6 with U6, but does interact with Lsm2-6p by two hybrid and immunoprécipitation assays (79). These data suggest that Lsm proteins form distinct complexes, one of which functions in decapping. Interestingly the Parker lab has demonstrated that the LSM1 complex affects decapping separate from the competition for cap binding between elF4E and Dcpip (77). Despite this, a protein that plays a role in translation initiation Mrtlp/Patlp/SpblOp is also needed for the LSM1 complex to stimulate efficient decapping. Mrti p, which is bound to the 40S subunit and polysomes, and is required for the association of the Lsm lp with polysomes (71).

The major exonuclease activity X m lp was also found to interact with the Lsm proteins through M rtip (78).

Xm l p requires a 5’ monophosphate at the 5’-terminus to degrade ssRNA or ssDNA generating mononucleotides (80). Following decapping the generation of a 5’-monophosphate on the 5’-terminus of the transcript produces a perfect product for Xm lp activity. Xmlp is the major activity for degradation of the decapped transcripts as XRN1 mutants accumulate deadenylated and decapped transcripts on the polysomes. This suggests that decay occurs in parallel with translation. In support of this, the interaction of Xm lp with the LSM1 complex through M rtip localizes the exonuclease to decapped polysome associated transcripts. Xm lp has high homology to Xrn2p, a nuclear protein that is required for RNA processing and is functionally interchangeable with X m lp (81).

17 The prevalent view today is that translation initiation factors play a role in regulating mRNA decay that is independent of their role in Initiation. This Is supported by the fact that a single amino acid change in elF5A leads to the accumulation of decapped mRNA (82), and that elF4E mutants have been shown to have a more rapid turnover of their mRNAs (83). Also a ts-elF4E has been shown to rescue the loss of decapping of the dcp1-1 mutant (77). Pabp inhibits decapping when bound to the poly(A) tail. This inhibition prevents decapping from occurring until deadenylation removes most of the poly(A) tail relieving Pabp inhibition of decapping. These observations imply that there may be a competition between the cap-binding complex and the decapping complex for binding to the ^'"G-cap. Transcripts that maintain the length of their poly(A) tail are stabilizing the interaction between Pabp and elF4F, which would maintain the competence for translation and protect the transcript from degradation.

However, if deadenylation proceeds, the interaction between Pabp and elF4F becomes compromised, decreasing translational efficiency, and thus making the cap structure more susceptible to binding and decapping by Dcpip leading to degradation of the transcript by Xm l p.

Work done in Roy Parkers lab has identified cis acting stability elements that account for the difference in stability between the stable PGK1 and unstable

MFA2 transcripts, which undergo degradation through this deadenylation- decapping pathway (84). MFA2 has been shown to have a cis instability element in the 3’UTR that is able to accelerate deadenylation and decapping of the

18 message. Interestingly, only when the context of the start codon was altered In the PGK1 transcript to create less efficient Initiation of translation did the rate of decay Increase. These data suggest that It Is possible for specific mRNAs to regulate their rate of decay through cIs acting factors even though they utilize the same degradation pathways (85).

1.4.2 3 -5' decay: the exosome.

A complex, called the exosome, which Is structurally very similar to the proteosome, exists in yeast and seems to be the major source of 3’-5’ exonuclease activity (86). The exosome Is composed of a core of at least ten proteins all of which have been proposed to be 3’-5’ exonucleases (see table

1.2). Six of these exonucleases appear to be 3’-5’ phosphorolytic enzymes since they are related to the E. coii3'-5' exonucleases RNase PH and PNPase.

Phosphorolytic exonucleases generate nucleotide diphosphates as their products. The other four proteins are thought to be hydrolytic enzymes that generate nucleotide monophosphates (72). Each protein Is approximately 30 kDa and the entire complex sediments In a glycerol gradient suggesting a size of

300-400 kDa. This Is consistent with there being only one copy of each protein present In the complex. Each of the subunits Is essential for the activity of the exosome since defects In one can affect the formation of the complex which Is essential for viability (86,87).

19 The exosome is an ATP dependent complex located in both the cytoplasm

and the nucleus. Like the proteosome there does not appear to be a substantial

pool of free exosome subunits. However, unlike the proteosome, the subunits do

retain their activity when they are not part of the complex. Also, homologous

cDNAs from humans can complement at least some of the phenotypes of yeast

with mutations in some of the exosome components (RRP4, RRP44, or CSL4

genes). This suggests that either the subunits are conserved to allow assembly

between different species, or more likely the subunits can function individually.

These facts suggest that the exosome subunits may have distinct functions

independent of the entire complex (72). The nuclear exosome has an extra

subunit, Rrp6p, which is also a 3’-5’ exonuclease (87). Rrp6p is the only subunit

of the exosome that is not essential for viability despite having strong defects in

all the known nuclear functions of the exosome .

Like the proteosome the exosome performs both processing and

degradation of specific substrates. Nuclear trimming of RNAs such as the 5.8S

RNA (86), as well as several snoRNAs and snRNAs (87), has been shown to be

reliant on the exosome. Since all of these functions require 3’-5’ exonucleolytic activity, it is likely that the exosome is the complex carrying out all of the processes. Mutations in some components of the exosome lead to accumulation of polyadenylated transcripts in the nucleus suggesting a role in nuclear RNA degradation (88). The exosome has also been shown to function in the degradation of pre-rRNA transcripts that are not properly processed (89). Other

2 0 possible functions would be the deadenylation of the telomerase RNA and

degradation of excised and debranched introns. Interestingly, the two major

functions of mRNA degradation and processing of rRNA can be genetically

separated. This is seen with the Ski4p (Csl4p) protein where the ski4-1 allele,

which contains a point mutation in the RNA binding domain strongly inhibits

exosome degradation of mRNA but has no effect on rRNA processing.

Conversely, the csl4-1 allele of the same gene affects rRNA processing but not

mRNA degradation (90).

Each function of the exosome identified to date is dependent on either

Mtr4p or Ski2p, which are closely related members of the superfamily II of RNA

helicases. This family of helicases interact with RNA and utilize ATP hydrolysis

to promote conformational changes in either RNA structure or RNA protein

Interactions (91). Defects in nuclear Mtr4p inhibit the processing of 5.8S rRNA,

snRNA, and snoRNA, as well as degradation of the 5’ extemal transcribed

spacer region of the pre-rRNA (91,87). Ski2p, which is found in the cytoplasm, is

required for the exosome to perform 3’-5’ mRNA degradation (92). It Is possible

that these helicases do not just function to unwind the RNA, they may also

function to target the exosome since many substrates that require these

helicases do not have any obvious secondary structure.

The PM-ScI complex in humans that was initially identified as an autoimmune antigen associated with polymyolytis-scleroderma overlap syndrome

2 1 has been demonstrated to have 9 of the 10 components of the yeast exosome.

The complex has also been shown to be present in both the cytoplasm and

nuclease, and it is functionally equivalent to the exosome. This suggests that the

exosome is conserved in higher eukaryotes.

1.5 mRNA decay in mammalian cells.

1.5.1 The role of AU-rlch elements (ARE) In mRNA decay.

Many unstable mammalian mRNAs contain regions that are rich in

adenosine and uridine. These regions are called AU-rich elements or AREs, and

are found in the 3’UTRs of many short lived mRNA species including cytokines

and proto-oncogenes. These unstable mRNAs have half-lives usually between

5-30 minutes. The ARE is the most widespread RNA stability determinant characterized in mammalian cells, and it is involved in the fine control of gene expression under many different physiological conditions (93). The ARE elements have been subdivided into 3 different subgroups, and it seems that they all require repeats of the pentamer AUUUA . It also seems that 5 to 6 pentamers are required in close proximity to each other for deadenylation (93).

The c-fos ARE is composed of two different regions. These two regions are Domain I, a 40 to 50 nucleotide region that contains several AUUUA pentamers, and Domain II, a U-rich region of about 20 nucleotides. If the sequence of 3 AUUUA pentamers is mutated the mRNA is stabilized five-fold

2 2 Factor Known function Source

DAN/PARN Poly(A) 3’-5’ exonuclease HeLa cells PMR-1 endonuclease serum protein/vitellogenin Xenopus liver mRNAs Xixernase endonuclease cleaves Xlhbox2B Xenopus oocytes G3BP phosphorylation dep. endonuclease (c-myc) Hamster Fibroblasts RNase L endonuclease MyoD/interferon-lnduced Myoblasts mRNAs vhs endonuclease host and viral mRNAs Herpes Simplex Virus ErEN endonuclease alpha-globin Erythroleukemia cells c-myc m RNase endonuclease c-myc Er^hroleukemia cells HL-60 endonuclease MONAP HL-60 cells mXrnIp 5’-3’ exonuclease (not dominate like yeast Murine Xrni p) hUpflp mRNA surveillance (ATPase/helicase) Human hUpf2p mRNA surveillance Human hUpf3p mRNA surveillance Human Exosome 3’-5’ exonuclease/ precise function still not Human known activitv? Known substrates Source

5-3’ exo uncapped mRNA Ascites cells 5’-3' exo/endo beta-globin w/nonsense codon MEL cells 3’-5’ exo IL-1 a Rab. Retie. Lysate 3’-5’ exo histone, poly(A) RNA Erythro. Leukemia cells 3’-5’ exo c-myc poly(A) tail and 3’UTR Erythro. Leukemia cells Endo Apo II Cockrel liver Endo IL-2 Jurkat cells Endo IGF-II Rat liver Endo Transferrin receptor Murine fibroblasts Endo? beta-tubulin Murine fibroblasts Endo Alu sequences in mRNA Erythro. Leukemia cells Endo beta-globin Rab. Retie. Lysate

Table 1.4 Major trans factors involved In vertebrate mRNA degradation.

23 while the rate of deadenylation is only slightly changed. However, if the U-rich region is deleted and the AUUUA pentamers are untouched, the deadenylation rate is decreased, and the message is stabilized two-fold (94,95). This example also demonstrates how the stability of an mRNA can have significant effects on cell physiology. The c-fos gene is only weakly onc:ogenic, but its oncogenicity increases approximately twenty-fold when the ARE is deleted (96,97,98,99).

The variety of ARE sequences suggests that there may be a number of different trans acting factors that interact with the elements. This imparts different regulatory characteristics on each group o f transcripts. Several different factors, such as AUF1, tristetraprolin, and the von Hippel-Lindau proteins have been demonstrated to destabilize mRNAs in a manner that is ARE dependant.

The ELAV protein HuR, on the other hand, has been shown to stabilize ARE containing mRNAs (100). A few of these factors will be elaborated on below.

AUF1/hnRNPD binding to an ARE correlated with the rapid decay of the mRNA both in vivo and in vitro (7). There are four altematively spliced variants of AUF1 (p37, p40, p42, p45) which are expressed] in most cells. The p37 and p42 display the most profound effect on mRNA stability (101). A.B. Shyu has suggested that individual mRNAs being bound by AUF1, which has been shown to shuttle between the nucleus and cytoplasm, may in part define the nuclear history and can influence its cytoplasmic fate (93). One example is the developmental regulation of phagocytes, which is a function of increased GIVI-

24 CSF expression due to a change in the half-life of the mRNA from 30 minutes in

neonatal cells to 100 minutes in adult cells (102). Immunodepletion of AUF1

causes this ARE containing message to be stabilized in neonatal cell extracts,

demonstrating the destabilizing effect of AUF-1 binding.

AUF1 can associate with itself to form dimeric and hexameric complexes, which bind the c-/bs ARE with nanomolar affinity to accelerate mRNA degradation (103). The formation of these multiprotein complexes is hypothesized to provide a large interacting surface for other proteins known to interact with AUF1, such as the complexes formed between AUF1 and the heat shock protein hsp70, translation initiation factor elF4G, or PABP (104).

Disruption of the interaction between AUF1 and elF4G allows for ubiquitination of

AUF1 and degradation by the proteosome. Heat shock induction of hsp70, down regulation of the ubiquitin-proteosome network, or inactivation of the ubiquitinating enzyme El all result in hsp70 sequestration of AUF1 in the perinuclease-nucleus, which leads to stabilization of the ARE containing mRNAs

(104). This allows for AUF1 to signal for the degradation of ARE containing mRNAs such as c-fos and quickly be removed from the system by the ubiquitin- proteosome pathway to regulate mRNA decay.

Another potential method to regulate ARE mediated degradation is signal transduction. One example of this is the regulation of IL-6 and IL-8 mRNA stability by a pathway that includes p38 and MAP kinase in transfected HeLa

25 cells and is dependent on the ARE in the 3’UTR of each message (105).

Another example of signal transduction pathways affecting ARE containing mRNA stability is the p-adrenergic receptor (P-AR) down regulation in human heart and murine smooth muscle cells (106). Stimulation of the P-AR by p- adrenergic increases AUF1 expression, which correlates with its binding to the p-AR mRNA 3’UTR and a 40-50% reduction of the mRNA. This likely accounts mechanistically for catecholamine-dependent desensitization of heart muscle cells and heart failure.

Recent work suggests that AUF1 binding to the ARE not only accelerate deadenylation, but also decapping. In HeLa SI 00 extracts, the rate of decapping can be inhibited 25-fold by the interaction between elF4E bound to the ^"^G-cap and the poly(A) tail (83). This supports the idea that a competition exists between the translation initiation complex and the decapping complex for binding to the 5’ termini of mRNA. ARE’s from TNF-a or GM-CSF mRNAs which are known to be bound by AUF1, stimulate decapping 7-10 fold. Formal proof of

AUF1 binding to the AREs in the in vitro decapping assay is still missing.

However, the data suggests AUF1 can not only accelerate deadenylation but also decapping.

Another class of ARE binding proteins are the embryonic lethal abnormal visual (ELAV) proteins. This class of proteins includes HuC, HuD, HuR, and Hel-

N1 (107). HuR which is present in both the nucleus and cytoplasm is expressed

26 in all proliferating cells while the other 3 ELAV proteins are neuronal specific.

Although HuR has not been shown to effect deadenylation it does delay degradation of the body of the mRNA. Binding of ELAV proteins at nanomolar affinity requires a minimum of 3 AUUUA pentamers which are, the same as those needed by AUF1 for a similar binding affinity (108). Given that HuR and AUF1 have opposite effects and bind the same region of the mRNA, they most likely act as antagonists with one another in regulating ARE-dependant decay of short­ lived mRNAs. Shyu has suggested that decay of ARE containing mRNAs is regulated by controlling the nucleo-cytoplasmic distribution of ELAV proteins in response to environmental stimuli. This would alter the ratio of ELAV to AUF1 allowing for fine control of mRNA stability (107).

Work done in the Steitz lab has identified 4 proteins that interact with HuR

(SETa, SETp. pp32, and acidic protein rich in leucine (APRIL)). They all bind to regions of HuR that are essential for its ability to shuttle between the nucleus and cytoplasm as well as stabilize mRNA. The nuclear retention of pp32 and APRIL by leptomycin B inhibition of CRM1 leads to increased association of HuR with these proteins as well as increased association with nuclear poly(A)+ RNA.

Furthermore, an increase in nuclear retention of c-fos transcripts is seen without any significant change in the distribution of total poly(A)+ RNA (109). These data suggest that the binding of these proteins affects the ability of HuR to bind some

ARE containing mRNA.

27 1.5.2 3 -5' exonucleolytic decay in mammalian cells.

Three different 3’-5’ exonucleases have been identified that play a role in

mRNA decay in mammalian cells (110,111,112). No prominent 5'-3’ exonuclease activity has been conclusively identified in mammalian cells despite the cloning of a murine homolog to Xm lp (mXmlp), which does not have the same dominant activity as the yeast X m lp (113). Although the predominate view

is that mammalian cells, like yeast, have a 5’-3’ degradation apparatus.

However, the inability to identify a predominate 5’-3’ activity prevents ruling out the possibility that 3’-5’ exonucleases could be the major form of degradation

after deadenylation. The major factors and activities that are believed to be

involved in mRNA degradation in vertebrates are listed in table 1.4.

One activity was identified in the murine erythroleukemia cell line K562 and is involved in c-myc mRNA degradation. The activity does not require ATP, is not Mg^"^ dependent, and does not require a ^"^G-cap to deadenylate an exogenous mRNA substrate in a manner that is reliant on the c-myc 3’UTR

(114). This nuclease produces 3’UTR degradation intermediates suggesting that it is not just a deadenylase. An in vitro RNA decay assay has been developed that produces the same c-myc degradation intermediates as found in vivo (114).

The second activity, a highly efficient 3’-5’ exonuclease, was identified in

HeLa cells and can be blocked by the poly(A) tail (111). This activity can also be blocked by a stable stem loop in the 3’ terminus of the message, or the poly(U)

2 8 tract that RNA polymerase III adds to the ends of its transcripts in a protein dependant manner. Importantly the presence of any of these structures further upstream does not block the activity suggesting that they act by prohibiting the loading of the exonuclease (115).

The third activity was identified by the Wilusz and Wahle groups. They have demonstrated that a 74 kDa protein, called poly(A) ribonuclease PARN (or

DAN), has deadenylase activity which is thought to represent the major deadenylase in mammalian cells (although no formal proof of this exists yet).

PARN has been shown to specifically interact with the 7'^G-cap structure and is stimulated by the presence of a poly(A) tail (116,117). Interestingly this interaction stimulates deadenylation. The Virtanen group has shown a 54 kDa fragment of PARN is associated with a 180-220 kDa oligomeric complex that prcessively degrades both capped and uncapped transcripts in vitro (118). This complex was also found to degrade capped transcripts 6 times better than uncapped transcripts. The 54 kDa fragment has been shown to be dependent on metal ions by the addition of aminoglycosides such as neomycin B which compete for binding to them (119). Since deadenylation is likely to be the rate limiting step in the initiation of degradation, the cap-PARN interaction is likely to be an important regulator of mRNA activity.

29 1.5.3 mRNA surveillance in mammalian cells.

Accurate mRNA synthesis is important for cell viability. One mechanism

that has evolved to degrade aberrant mRNAs is called mRNA surveillance or

nonsense mediated decay (NMD). mRNA surveillance removes transcripts that

contain premature stop codons, retained introns, skipped exons, and extended 3'

or 5’ UTRs all of which can negatively impact on the stability or translation of a

message. Recently mRNA surveillance has been shown to also be important for

the decay of ‘normal’ mRNAs such as those containing uORF’s or altematively

spliced messages (120). One example of this regulation is AUF1 mRNA which

has 4 altematively spliced 3’ UTR variants, discussed later.

This mode of decay shares much of the degradation machinery already

mentioned with some notable differences in the mechanism of decay. In

contrast, standard mRNA decay proceeds by slow deadenylation then decapping

and degradation by 5’-3’ exonucleases; NMD can decap a message independent

of deadenylation. In yeast, this is accomplished by Dcpip (120). This pathway

requires the UPF proteins (Upflp, Upf2p, and Upf3p), prior translation of the

message, and formation of the surveillance complex as described below.

Decapping doesn’t require the LSM-1 complex or Mrtip like the default pathway

suggesting that the decapping enzymes are recruited by another mechanism

(71). The surveillance complex, which consists of Upflp, Upf2p, Upf3p, and the translation release factors eRFI and eRF3, acts to scan the mRNA to determine if the transcript is “proper” or should be degraded (121).

30 In Saccharomyces cerevisiae, during translation termination the local mRNP environment may be ‘sensed’ by the surveillance complex. If the context is poor the complex scans the downstream sequence. The identification of a downstream element (DSE) will dictate that nonsense mediated decay (NMD) will occur (122). The DSE’s that have been identified to activate NMD are typically degenerate sequences that may be present in multiple copies in a mRNA.

There are two theories as to how this works. One is that at termination the surveillance complex scans downstream, and if a DSE is identified NMD will occur. Otherwise, either re-initiation, if there is a uORF, or termination will occur.

The second theory is that DSE’s influence how translation termination occurs.

This model considers factors such as the architecture of the mRNP, a Upfl p timing mechanism, and a restructuring of the mRNP to a form that promotes stability following translation termination (120).

Upfl has been demonstrated to function in both NMD and translational termination along with eRFI and eRF3 suggesting just how intimately these processes are tied together. The potential role of Upfl p as a kinetic clock in determining the difference between proper and improper termination is suggested by the fact that local and distal sequence elements along with associated factors can influence the context of the formation event of the translation termination process (123). For example if termination is too slow or

31 improper Upfl p, a RNA helicase, and its ATP co-factor will cause a structural rearrangement of the terminating altering its fate (120).

Work done by the Peltz lab has identified a protein that binds to the DSE,

Hrpip (124). Mutation of the Hrpip gene stabilizes messages degraded by NMD without affecting the decay of normal transcripts. Hrpip has been shown to bind with Upfl p while bound to the DSE, while the mutant form can not bind the DSE or interact with U pflp, suggesting that it is the factor that binds to the DSE in yeast and signals for NMD to occur.

The three major yeast proteins Upfl p, Upf2p, and UpfSp have all been found to have a human homolog (hUpfl, hUpf2, hUpfS), and hUpf2p and hUpfSp have been demonstrated to function in NMD (125,126). Despite this conservation of factors there do appear to be some key differences in the mechanism of identification of substrates for decay between yeast, described above, and mammalian cells.

The key element in identifying substrates for NMD in mammalian cells is the presence of a splice junction downstream of the termination codon. This suggests that splice sites are somehow marked in a way that is present on the translated mRNAs. The Moore and Maquat groups have suggested that Rnpsi,

PrpSp, and SrmlSO are all possible candidates to mark the splice junctions (127).

The Steitz group has shown that hUpf2 and hUpfS when bound downstream of

32 the termination codon via fusion to the MS2 coat protein can lead to NMD. They

hypothesize that they may bind proximally to splice junctions and are recognized

by the surveillance complex, if they are located downstream of the stop codon

identifying premature termination and signaling for NMD (126).

One example of how this mechanism is used to degrade ‘normal’ mRNA is

in the degradation of the 4 altematively spliced AUF1 mRNA including one that

contains a 107 nt exon (exon 9) and the adjacent downstream intron 9. Work

done by the Peltz and Brewer groups has shown that intron 9 can regulate

stability by two mechanisms. The first is two AUF1 binding sites that when

deleted from an unspliceable intron stabilize the message. Interestingly, if deleted from a spliceable intron 9 there is no effect on the stability of the message. However, if the spliceable intron with the deleted AUF1 binding sites is co-transfected with a dominate negative UPF1 the destabilization is blocked.

This suggests that NMD is the method of decay for the wild type mRNA.

In yeast a nuclear degradation pathway exists that removes defective pre mRNAs using the exosome and Rati p/Xm2p a 5’-3’ exonuclease. In vertebrate cells, aberrant pre-mRNAs that have not been spliced properly or are otherwise deficient in mRNP structure are rapidly degraded. However, even though the exosome complex has a homologous complex identified in vertebrate cells and a murine X m lp has been identified, it is not known if this pathway is conserved in humans.

33 NMD bypasses two key control systems for the regulation of mRNA stability. The first is that the poly(A) tail does not have to be shortened to oligo A lengths prior to decapping. The second being features of the mRNA itself that have been found to affect the rate of decapping. For example, the mRNA for

PGK1 is normally stable following deadenylation due to a slow rate of decapping.

However, if a nonsense codon is introduced to initiate NMD it is rapidly decapped. This suggests that NMD does more than Just nullify the stabilizing effect of the poly(A) tail probably triggering additional alterations in mRNP structure that specifically promote decapping (120).

1.6 mRNA endonucleases in vertebrate cells.

Evidence suggests that endonucleases may play a more significant role in mRNA degradation in vertebrate cells than expected. A number of labs have identified specific cleavage products in a number of different systems. However, only a few labs have been able to identify the enzyme that is responsible for these cleavages (128,129,49). Mapping of these potential endonuclease cleavage sites suggest many endonucleases have a sequence selectivity for a specific motif such as AU-rich elements (130), or a preference for a specific sequence in a specific structural context such as the transferrin receptor (TfR) iron response element (IRE) (131). However, most of the cleavage products identified have suggested that this specificity is loose, and multiple cleavages are often possible for a given enzyme activity. Many of these activities have been

34 identified as associated with the translational apparatus, suggesting either the localization of the transcript to an endonuclease associated with the ribosome, or that the rearrangement of the mRNP structure may be needed for cleavage

(132,133). Although polysome association is important for the destabilization of a number of transcripts, Xlhbox2B mRNA in Xenopus, and a-globin in murine erythroleukemia (MEL) cells demonstrate that not all transcripts are dependent on polysome association to be destabilized (128,6).

From the number of systems in which endonuclease cleavages initiate decay of a transcript, one of the best described is the transferrin receptor mRNA.

When intracellular iron levels are low, the iron response element binding protein

(IRE-BP) binds to one of five IRE stem loops in the 3’UTR, occluding an endonuclease cleavage site, thereby stabilizing the transcript. When iron level within the cells increase, IRE-BP dissociates from the IRE leading to endonuclease cleavage and degradation of the transcript (134). The avian apolipoprotein II (apo II) mRNA is induced and stabilized by estrogen. When estrogen is withdrawn, the transcript becomes unstable. The endonuclease activity has been shown to have a preference for cleavage in 5’-AAU-3’ or 5’-

UAA-3’ sequence motifs present in an unpaired loop region in the 3’UTR (135).

An erythroid-enriched endoribonuclease (ErEN) activity has been identified that cleaves within the a-complex in the 3’UTR (6). Binding of PABP to the poly(A) tail stabilizes complex binding to the 3’UTR which in tum stabilizes the mRNA. After deadenylation, PABP can no longer bind leading to dissociation

35 of the a-complex and destabilization of the mRNA by endonuclease cleavage (9).

G3BP, a 120 kDa Ras GAP src homology 3 binding protein, binds to the GTPase activating protein, RasGAP, at the plasma membrane of serum stimulated, but not quiescent Chinese hamster lung fibroblasts. It has a phosphorylation dependent endonuclease activity that cleaves c-myc mRNA in the 3’UTR. Serine hyperphosphorylation in quiescent cells activates the endonuclease, and this links mRNA decay with the ras signaling pathway (49).

Some mRNAs are regulated by multiple pathways that utilize different endonucleases. The best example of this is the c-myc mRNA. As mentioned above, it can be regulated by G3BP cleavage within the 3’UTR. However two other activities have also been identified that regulate c-myc mRNA. First, an

ARE element has been identified in the 3’UTR of c-myc. Second, the Ross lab has identified a coding region determinate (CRD) in the 3’ portion of the coding region that has an endonuclease cleavage site. The CRD is targeted by a polysome associated endonuclease. Cleavage is dependent on ribosome pausing within the first 70 bp of the CRD due to 1 or 2 rare codons. This renders the CRD susceptible to cleavage. Mutation of these sites to more common codons prevents pausing and stabilizes the message. A binding protein (CRD-

BP) has also been identified that can bind to the CRD and protect it from endonuclease cleavage (50).

36 1.7 Estrogen regulation of mRNA stability.

A number of steroids have been demonstrated to regulate mRNA stability.

Estrogen is the best studied of these (136,137,138). Work done by the Shapiro

group has demonstrated that vitellogenin mRNA is stabilized by estrogen in

Xenopus liver on polysomes mediated by vigilin, a 155 kDa estrogen induced

protein that binds the 3’UTR in a unstructured single stranded region. The 14 KH domains are hypothesized to wrap the RNA around the protein and stabilize the mRNA by preventing polysomal ribonuclease (PMR-1) cleavage within this

region at two consensus cleavage sites (41,139).

The Williams group demonstrated that apolipoprotein II (apo II) translation is induced by estrogen, and following withdrawal, the mRNA is rapidly degraded by an endonuclease activity that cleaves a 3’UTR stem loop structure in the loop domain with the sequence 5’-AAU-3’ or 5’-UAA-3’ (135). Eight estrogen induced proteins were identified that bind to the apo 11 mRNA. Only one of these was found to lose binding when the cleavage site was removed. However it is not know whether this 58 kDa protein is the endonuclease or a protecting factor (30).

In MCF-7 cells, estrogen has been shown to regulate the stability of ER-a.

Estrogen causes a 6 -fold decrease in the stability of the ER-a mRNA from 4 hours to 40 minutes (140,141). Interestingly, pactamycin and puromycin (both inhibit polysome association with mRNA) abrogated the effect, while cycloheximide (inhibits elongation) didn’t have any effect on the destabilization of

37 the mRNA. This suggests that the ER-a mRNA is destabilized by a ribosome

associated nuclease. It is not yet known if this is an endo or exonuclease.

1.8 identification and characterization of PMR-1.

In Xenopus liver, estrogen causes induction of transcription and

stabilization of the vitellogenin transcript (142,143,144). May et al. have also

demonstrated that 95% of the albumin mRNA disappears after 4 days of

estrogen stimulation and retums to normal levels over the next 12 days (145). It

was also demonstrated that transcriptional levels did not change in liver cube

culture suggesting that it is a post- transcriptional effect that leads to these

changes in albumin levels (146). Further work demonstrated that there was a 3-

fold destabilization of the albumin mRNA caused by estrogen in male and female

Xenopus primary hepatocyte cell culture. It was also observed that

concentrations of albumin were higher in male frogs than in female frogs.

Hormonally dependent stability effects were more pronounced in the male cells.

This was believed to be due to the estrogen background that was present in the

female cells (147).

Riegel et al. determined that 12 hours after estrogen induction estrogen

causes albumin mRNA to drop from 30,000 molecules per cell to 7,000

molecules per cell. No decrease is detected until 10 hours after induction, and

levels of an unrelated non-serum protein mRNA, p-globin, did not change (148).

This was then demonstrated to be ER-dependent (149). Estrogen was

38 demonstrated to have no effect on splicing or polyadenylation site selection in the

3’UTR of the albumin mRNA (150). Also, no change in the transcriptional initiation site or state of albumin capping was identified. When the extranuclear fraction of primary hepatocytes was examined, a >90% decrease in albumin mRNA levels was seen between 9 and 12 hours. Another observation was that both the 74 and 6 8 kDa isoforms posses a short 17 nucleotide poly(A) tail. This suggests that unlike many mRNA that require prior deadenylation as a rate limiting step before decay of the message, both forms of albumin already possess a short poly(A) tail. This does not mean that the 3’ portion of albumin is not providing a stabilizing effect similar to transcripts with a long tail, but it does suggest that degradation is independent of deadenylation.

Work done by Pastori et al. demonstrated that estrogen regulates more than just albumin mRNA stability. Other serum protein mRNAs are also destabilized by estrogen such as y-fibrinogen, transferrin, inter-a-trypsin inhibitor, and serum protein 12B (151,152). y-fibrinogen, like albumin, was demonstrated to disappear by a post transcriptional ER-dependant manner (153). Sucrose gradients showed that serum protein mRNAs disappeared with similar kinetics from both mRNPs and polysomes, and that all the above mentioned transcripts,

-85% of the secreted serum mRNA in Xenopus liver were likely degraded by the same mechanism (151). The degradation of all these mRNAs was shown to occur in the cytosol, which suggested that it was regulated by a ribonuclease(154).

39 Subcellular fractionation showed that 75% of the activity was present on the polysomes and only about 10% was present on the mRNPs. Polysome fractions when incubated with a [^^P]-labelled albumin transcript showed a preference for degradation of albumin over feritin by an estrogen stimulated activity. This was believed to be an endonuclease, since it produced a discrete band. Early sucrose gradients following PMR-1 by activity assay suggested that

PMR-1 associates with the 80s and polysomes and not the individual subunits (132). However, later studies using antibodies, as well as activity, demonstrated that PMR-1 is associated with both the polysomes and the mRNPs. Estrogen stimulation leads to a 22-fold increase in the unit activity of

PMR-1 on polysomes while no significant increase in the unit activity of PMR-1 associated with the mRNPs was seen (155).

The in vitro production of discrete degradation products was used to purify the enzyme responsible for the ER-dependent destabilization of these serum protein mRNAs. The purified activity was identified as a 62/64 kDa doublet by

SDS-PAGE. Isoelectric focusing showed that the protein was basic and separated into 3 dots at pi 9.6, 9.7, and 9.9 (129). The activity was found to be independent of divalent cations, resistant to placental ribonuclease inhibitor, and does not require a RNA . Experiments using a TAP-tagged PMR-1 have demonstrated that PMR-60 is not tyrosine phosphorylated when expressed in mammalian cells. This is consistent with data using PMR-1 (Yang, P., and

Schoenberg, D.R., personal communication). In vitro mapping showed a

40 preference for the sequence APyUGA, although It was able to cleave the

message at other sites as well. The sequence APyUGA is present 14 times in

albumin, 9 times in transferrin, and 7 times in y-fibrinogen. However, it is not

present in the intercellular ferritin protein mRNA which is not specifically

degraded by this activity.

Tryptic digest of the purified protein generated 5 peptides that were

sequenced, however the N-terminus was blocked and therefore unable to be

sequenced. Degenerate primers designed from these peptides were used to

clone PMR-1 from a Xenopus liver cDNA library (156). The 2512 bp cDNA

encoded a 80 kDa polypeptide that is processed to an amino-terminal truncated

62/64 kDa form. This was found to have no sequence similarity to the

superfamily of vertebrate endonucleases. Instead, a 57% sequence identity was

found with human myeloperoxidase (hMPO).

1.9 Human myeloperoxidase.

Human myeloperoxidase is an antibacterial agent that is found only in the

azurophilic granules (or primary lysosomes) of human polymorphonuclear

leukocytes (157). It is part of the host response to bacterial infection generating

hypochlorous acid in the presence of hydrogen peroxide and chloride ions. It consists of a 120-150 kDa heterotetramer composed of two 55 kDa heavy chains and two 15 kDa light chains. Both of these chains are encoded on the same mRNA and assembled into the macromolecular complex following proteolytic

41 processing of the precursor protein. This processing occurs in the Golgi concomitantly with the addition of both heme and N-linked carbohydrates (158).

Due to the presence of a novel covalent sulfonium ion linkage between the vinyl group on pyrrole A ring and methionine 409, hMPO has a characteristic green color (159). Two histidines (His^®\ His^°^) have been identified by site directed mutagenesis to be involved in the coordination of the iron group within the heme.

Mutation of either to an alanine leads to a loss of peroxidase activity (160,161).

1.10 Purpose of this study.

This dissertation sought to address three issues:

1. Characterize the biochemical differences of the related frog polysomal

ribonuclease and human myeloperoxidase.

2. Identify PMR-1 expression in Xenopus and murine tissues as well as cell

culture, and address when PMR-1 is phosphorylated in Xenopus liver.

3. Develop a sensitive method to identify degradation intermediates of in vivo

mRNA decay.

1.10.1 Biochemical comparison of PMR-1 and hMPO.

Since the Xenopus endonuclease PMR-1 was found to have a 57% identity with human myeloperoxidase, it was important to define whether PMR-1 is frog MPO or just a closely related protein. This was accomplished by identifying the biochemical similarities and differences between these two proteins. The following were used to define the relatedness of these two

42 proteins, the presence or absence of heme, N-linked glycosylation, and autolytic

cleavage of the peptide backbone upon heating. Lastly, the enzymatic activity of the two proteins was compared. These comparisons will help define the

relationship between these two similar proteins.

1.10.2 Expression of PMR-1 in Xenopus and murine tissues and estrogen

phosphorylation of PMR-1.

It is generally believed that there are a small number of endonucleases

that are used to regulate the stability of a large number of transcripts in a tissue

and environment specific means. To address the possibility that PMR-1 may

have a more diverse function than just the estrogen regulation of serum protein

mRNAs in Xenopus liver, we looked for expression of PMR-1 in other Xenopus tissues as well as mouse tissues and human cells in culture.

Estrogen activates PMR-1 by a mechanism that does not require any new transcription. This suggests that activation of PMR-1 is most likely to occur by phosphorylation of the protein. However, PMR-1 was shown to not be tyrosine phosphorylated, suggesting that PMR - 1 is not activated by tyrosine phosphorylation (Yang,P., and Schoenberg,D.R., personal communication). We wished to look at general changes in phosphorylation of PMR-1 with Calf

Intestinal Alkaline Phosphatase (GIAP), to see if estrogen had any effect on the phosphorylation of PMR-1.

43 1.10.3 In vivo mapping of endonuclease cleavage sites.

With the exception of insulin-like growth factor II mRNA, whose unique structure results in a remarkably stable in vivo endonuclease degradation product

(162), most mRNAs are degraded without significant accumulation of decay intermediates. However, decay intermediates have been observed using crude in vitro decay systems. The most likely explanation of this is that in vivo these intermediates are rapidly degraded by exonucleases like the products of RNase

E degradation in prokaryotes. A few labs have mapped in vivo cleavage sites using primer extension or SI nuclease protection assays (Fig 1.1)(163,134).

However, these methods are not very efficient for this purpose and can require over 30 day exposures making it very difficult to differentiate endonuclease cleavage sites from polymerase pause sites. This led to a need for a more sensitive assay that could be used to identify the 3’ ends of in vivo degradation intermediates produced either by endonuclease cleavage or pausing of a 3’-5’ exonuclease.

This was accomplished by designing a ligation mediated polymerase chain reaction (LM-PCR). Briefly, a primer bearing a 5’-phosphate and 3’-amino group is ligated onto a population of total cellular RNA. This sen/es as the template for reverse transcriptase primed within the sequence of the ligated primer. PCR is then performed using a 5’-[^^P]labeled primer specific to the

44 Figure 1.1 Identification ofin vivo cleavage products by primer extension. A. Ten |ig of total liver RNA from control (lane 6 ) or 12 hr estrogen-treated frogs (lane 7) was analyzed by primer extension using an end-labeled primer to position 280-301 of the albumin mRNA. Twenty ng of albumin 5’ transcript was also examined as a control for primer extension stop sites (lane 5). The 7 unique primer extension products detected in RNA from estrogen-treated frogs are indicated by the numbers on the right side of the gel, which correspond to the LM-PCR bands (Fig 4) that mapped to the same site. B. The primer extension products are shown graphically on a secondary structure model for wild type RNA. Filled arrows correspond to the cleavage sites that were previously mapped in vitro. 0. Ten pg of total liver RNA from 12 hr estrogen-treated frogs was analyzed by primer extension following incubation at 22°C for 30 min with no additions (lane 5 and 7) or with the addition of 100 units (lane 6 ) or 200 units (lane 8 ) of purified PMR-1. The position of the unique primer extension products are indicated by the numbers on the right side of the gel, which correspond to the LM-PCR bands (Fig 3.16) that were mapped to the same site. In each experiment a DNA sequencing ladder from the same primer was used to determine the location of the unique primer extension products on albumin mRNA.

45 PMR-1 A T G C « ^ C M + E

11.

180 -

c c

AO

in VIVO in vitro

Figure 1.1 Identification of in vivo cleavage products by primer extension.

46 mRNA of interest and the primer used for reverse transcription. The PCR

products are separated on a denaturing polyacrylamide/urea gel, excised, re­ amplified and sequenced. The junction of the ligated primer identifies the 3’ end of a particular degradation intermediate.

47 CHAPTER 2

MATERIALS AND METHODS

2.1 Purification of PMR-1 from Xenopus liver.

The following protocol was adapted from (129) and (154) for 30 adult male frogs which generate between 75 and 100 g of liver. Frogs are injected with 1 mg of 17|3-estradiol in 0.1 ml of propylene glycol/DMSO (9:1, v/v) 9-48 hours prior to harvesting the livers. Then anesthetized with 3-aminobenzoic acid ethylester (Tricane) for 30 minutes and the livers are removed using standard sterile tools and techniques. It is important to remove the gall bladder without rupturing because of all the proteases that are present.

2.2 Crude liver polysome isolation.

Tissues are perfused with 1 X SSC, blotted dry and weighed, then chopped into 1 mm cubes and 2.5 ml/g of homogenization buffer (40 mM Tris-

HCI, pH 7.5, 10 mM MgCIa, 7% sucrose, 2 mM DTT, 0.2 mM PMSF, 0.5 pg/ml leupeptin, 0.7 pg/ml pepstatin A, 2 pg/ml ap rotin in, 50 mM NaF, 0.2 mM NaVOa) was added. Tissue is homogenized with a Wheaton Overhead Stirrer and tapered tissue grinder with Teflon® pestle at a speed of 4000 RPM for two minutes at 4 °C. Homogenized lysate is centrifuged at 1000 x g for 10 minutes at

48 4 °C to remove large nuclei and insoluble material and then further centrifuged at

25, 000 X g for 15 minutes to remove all subcellular organelles. The post-

mitochondrial supernatant is centrifuged at 1 0 0 , 0 0 0 x g for one hour to pellet the

polysomal material.

The pellet is dissolved in one-half the volume of homogenization buffer

with 50 mM EDTA and 500 mM KCI. Extracts are rocked overnight at 4 °C to

extract PMR-1 from the polysomes. The sample is then centrifuged at 206, 000 x

g for 1 hour, this will pellet the ribosomal subunits and leave PMR-1 in the

supernatant. The polysomal salt extract is then concentrated into a volume of

less than 20 ml for the subsequent chromatography steps. This is accomplished

by centrifugation through a Millipore concentrator, then samples are dialyzed into

40 mM Tris-HCI, pH 7.5, 50 mM NaCI, 2mm DTT, 0.2 mM PMSF, 50 mM NaF

and 0.2 mM NaVOa overnight at 4°C.

2.3 FPLC purification of PMR-1.

The following chromatography protocol utilizes the Pharmacia (FPLC)

system consisting of a P-500 High Precision Pump, a GP-250 PLUS Gradient

Programmer, a FRAC-200 fraction collector and a FRAG-2 Chart Recorder.

Chromatography first involved passage of the polysomal salt wash from section

2 . 2 through a series of 3 Econo-Pac Q (QAE) cartridges (5 ml, Bio-Rad) to

remove negatively charged proteins. PMR-1 will not bind the cationic matrix.

The flow-though is dialyzed against a phosphate buffer (50 mM sodium 49 phosphate, pH 6.75, 2 mM DTT and 0.2 mM PMSF) overnight and is then applied to a series of 2 Econo-Pac S (SE) cartridges (5 ml, Bio-Rad). PMR-1 does bind this anionic column and was eluted with a gradient of NaCI (0.01-0.5

M) with the protein eluting between 200 and 350 mM. Fractions are collected and assayed for PMR-1 activity in vitro using a uniformly radiolabeled [a^^P-

UTP]-albumin RNA (Xa470, see section 2.11.2) to determine which fractions have PMR-1 activity. These fractions are pooled and dialyzed against 50 mM sodium phosphate, pH 6.75, 2 mM DTT and 0.2 mM PMSF ovemight. The samples are applied to a single 5 ml column of Econo-Pac CHT-II

(hydroxyapatite-Bio-Rad) and eluted with a linear gradient of 0.01-0.5 M NaCI.

Fractions are again assayed in vitro to determine which possessed PMR-1 activity. These final fractions are pooled, dialyzed against 40 mM Tris-HCI, pH7.5, 2 mM dithiothreitol, 20% (v/v) glycerol, 0.2 mM PMSF, aliquoted and stored at -80°C until needed.

Samples are evaluated by SDS-PAGE and silver staining to determine their purity. Protein concentrations are determined by Bradford assay, see section 2.3.2. A typical purification generates approximately 12.5 fig of protein or

5000 units of PMR-1 (Where one unit is been defined as the amount of PMR-1 that completely degrades 7 fmol of albumin transcript in 30 minutes at 22 °C), with an approximate 235-fold purification of PMR-1.

50 2.4 Bradford assays.

Bradford reagent (Bio-Rad, Hercules, CA) Is diluted 1:4 with water and filtered through a Whatman number 1 filter. Reactions are carried out In a 96 well dish with 200 |ii of diluted reagent added to each well. Protein diluted with water to 10|il Is added and Incubated at room temperature for 10 minutes before the absorbance at 595 nm Is determined using a Beckman DU 640 spectrophotometer. Either IgG or BSA of a known concentration Is diluted in the sample buffer and used to derive the standard curve for each assay

2.5 Autolytic cleavage of hMPO.

The ability of hMPO and PMR-1 to autolytlcly cleave themselves Is tested by following a modified protocol from Taylor et al. (164). Samples of either 1 pg hMPO or 10 pg polysomal liver extract are Incubated at room temperature for 15 minutes, add 5% p-mercaptoethanol to one third of them. All samples are then

Incubated at 95 °C for 8 minutes. After cooling the samples add 5% (3- mercaptoethanol to another third. The samples are them western blotted as described In section 2.5 using a polyclonal antibody raised against either hMPO purchased from Dako (Carplnterla, CA), or PMR-1 described In (156)

2.6 Western blotting.

Protein samples are prepared In an equal volume In IX SDS sample buffer (50 mM trIs-HCI, pH 6 .8 , 50 mM |3-ME, 2% SDS, 0.1% (w/v) bromophenol blue, 10% glycerol) then heated to 1 00 °C for 5 minutes. Samples are then 51 electrophoresed on a 10% polyacrylamide gel (37.5:1 acryllmideibis-acrylamlde) at 150 V until the bromophenol blue dye has migrated to the bottom of the gel.

This is then electroblotted to a PVDF membrane that has been presoaked in

100% methanol, and then western transfer buffer (39 mM Glycine, 48 mM Tris- base, pH 8.3, 1.35 mM SDS and 20% methanol (v/v) for either 90 minutes at 200 mA, or 16 hours at 15 mA). The PVDF membrane is then blocked in a solution of

TEST (20 mM Tris-HCI, pH 7.5, 150 mM NaCI, 0.5% Tween-20) with 5% (w/v) non-fat dried milk (carnation) for 2-16 hours at 4 °C. The membrane is then probed with a primary ( 1 °) antibody for one hour at room temperature with the titer ranging from 1:1000 to 1:10,000 (v/v) in TEST plus 5% milk. Then the blot is washed 3X, 1 0 minutes each, with 50 ml of TEST. The secondary ( 2 °) antibody is then incubated for one hour with a titer between 1:10,000 to 1:30,000 in TEST plus 5% milk. Again the blot is washed 3X, 10 minutes each, with 50 ml of TEST.

After a final wash withi X TES the protein is visualized using chemi- luminescence

2.7 Spectrum of hMPO and PMR-1.

To determine if there is a heme group present 0.1 pg hMPO or 0.02 pg of purified PMR-1 (Dompencial et al.) in 50 pi of buffer (40mM Tris, pH 7.5, 2 mM

DTT) is used to perform a wavelength scan from 200 nm to 600 nm using a

Eeckman DU 640 spectrophotometer.

52 2.8 Endoglycosidase H cleavage

1 |jg hMPO and 10 pg crude polysomal extract are incubated at 100 °C for 10 min with denaturing buffer (5% SDS, 10% p-mercaptoethanol). They are allowed to cool to room temperature and then G5 buffer (0.05 M Sodium Citrate pH 5.5) and 750 units of endoglycosidase H from New England Biolabs (Beverly,

MA) are added, and incubated at 37 °C for 1 hour. Each reaction is then electrophoresed on a 10% SDS-PAGE gel. Which is then transferred to an

Immobilon P membrane and Western blotted as described in section 2.5, using a polyclonal antibody raised against either hMPO purchased from Dako, or PMR- 1 .

2.9 Concanavalin A binding.

40 pi of Concanavalin A linked to Sepharose 4B beads both purchased from Amersham Pharmacia Biotech (Piscataway, NJ) is equilibrated with 400 pi of wash buffer (20 mM Tris, pH 7.5, 500 mM NaCI, 1 mM MgCI, 1 mM CaCI), with gently rocking for 1 hour at 4 °C. The Beads are pelleted by centrifugation at

1.000 X g for 30 seconds. The supernatant is removed and 400 pi of buffer with either 0.5 pg of hMPO or 2 pg of a partially purified PMR-1 extract is added. The supernatant is removed and the beads are washed four times with 4 volumes of wash buffer. Bound protein is then removed by three more washes with elution buffer (20 mM Tris, pH 7.5, 500 mM NaCI, 1 mM MgCI, 1 mM CaCI, 500 mM methyl-a-D-mannopyroside). Each sample is precipitated by adding 1 / 1 0 the volume of TCA and incubating on ice for 20 minutes. They were centrifuged at

13.000 X g for 15 minutes and the pellets are dissolved in 20 pi of 0.5 M Tris pH

53 8 .8 , 0.4% SDS. Western blots are performed as described in section 2.5 and probed with either a PMR-1 polyclonal antibody or a hMPO Monoclonal antibody purchased from Dako.

2.10 hMPO ribonuclease activity

hMPO is assayed to determine if it possessed an endonuclease activity similar to that of PMR-1, by incubating 1 pg, 2 pg, or 3 pg of pure hMPO along with a fragment of uniformly labeled [a-^^P-UTP] albumin mRNA transcript in a 40 mM Tris-CI pH 7.5, 2 mM DTT buffer for 30 minutes at 22 °C. These same reactions are also performed in the presence of 1 mM Mg^^ since many ribonucleases are activated, or have there activity enhanced, by Mg^^ including

PMR-1. Samples are electrophoresed on a 6 % urea/acrylamide gel, and ribonuclease activity is assessed by drying the gel and subjected it to autoradiography to visualize the products.

2.11 Peroxidase activity assay.

Peroxidase activity is assayed using 2,2’-azinodi(3-ethyl- benzthiozoline sulphonate) or ABTS from Amresco. One hundred nanograms of hMPO or PMR- 1 purified from liver polysomes, described in section 2.3, is added to 200 pi of 1 mM ABTS in 50 mM sodium citrate, 150 mM sodium phosphate, pH 4.4, containing 0.01% H 2 O2 . Incubated for 15 minutes at 25 °C, following which the absorbance is measured at 405 nM using a Beckman DU 640 spectrophotometer.

54 2.12 In vitro activity assays

A 5’[^^P]labeled transcript corresponding to nt 1690-2002 of the albumin mRNA is prepared as described previously (163). This is incubated with protein extract in a buffer containing 30 mM Tris-HCI, ph 7.5, 1 mM DTT, 2 mM MgCIa and 75 mM KCI, at 25°C using 10 pg of unfractionated polysome extract from estrogen-treated frogs. The reactions are stopped between 2-60 minutes latter by adding one volume of stop solution (98% formamide (v/v), 0.1% (v/v) bromophenol blue and 0 .1 % xylene cyanole) and heating at 95 °C for 3 minutes.

The samples are then electrophoresed on a 6 % polyacrylamide/urea gel, and cleavage products are visualized by autoradiography.

2.13 Purification of liver RNA: guanidinium-isothiocyanate method.

All steps should be performed at 4 °C. This method for RNA extraction is used for liver tissue samples. Liver samples are excised as described in section

2.1 and chopped into small cubes. Guanidine solution is added (10 ml/g tissue,

5M guanidine isothiocyanate, 50 mM Tris-HCI, pH 7.5, 10 mM EDTA, 735 mM p-

ME) and the mixture is homogenized using a Wheaton Overhead Stirrer and tapered tissue grinder with Teflon® pestle at a speed of 4000 RPM for two minutes at 4 °C. The homogenate is centrifuged at 12, 000 x g for 1 0 minutes at

12 °C to remove any insoluble material. One tenth volume of 20% (w/v) N- lauroylsarcosine is added and heated in a water bath at 65°C for 2 minutes to denature the protein. 0.1 g CsCI/ml of liver extract is then added, it is important to be certain that the CsCI is completely dissolved before proceeding to the next 55 step. The extract is layered over 9 ml of 5.7 M CsCI in a sialinized, polyallomer

tube and centrifuge ovemight at 113, 000 x g at 22 °C in a Sorvall TH-641

swinging-bucket rotor.

The next day, the supernatant is carefully removed by placing the end of a

Pasteur pipette on the top of the extract, removing the extract from the top down.

There should be a white band just below the midpoint of the gradient that

represents sheared DNA; care should be taken to remove all of this to prevent

contamination of the RNA present as a clear pellet at the bottom of the tube.

Once all of the supematant has been removed, the tube is inverted for 10

minutes to drain away any remaining liquid. 3 ml of resuspension solution (5 mM

EDTA, 0.5% (w/v) N-lauroylsarcosine, 5% (v/v) (3-ME) is added and the bottom

and sides of tube are washed using a pipette. The tube is incubated at 4 °C for

24 hours to resuspend the RNA. The high concentrations of (3-ME and N-

lauroylsarcosine will prevent degradation during this step.

Following resuspension, the RNA is extracted once with one volume of phenol:chloroform;isoamyl alcohol (25:24:1) and once with one volume of chlorofoim.isoamyl alcohol (24:1). Sodium acetate, pH 5.2 is added to 0.3 M and

2.5 volumes of ice-cold ethanol is also added and the RNA is precipitated 0/N at

-20°C. The RNA is washed with 70% ethanol and the pellet is dissolved in an appropriate buffer, such as water or 1 0 mM Tris-HCI, pH 8 .0 , just before use.

56 2.14 Purification of total RNAfrom cell culture using TRIzol®.

TRIzol® reagent is a phénol based solution used to extract nucleic acid from tissue cultured cells in one step. The following is a modification of the manufacturers protocol designed to produce cleaner RNA. 2 ml of TRIzol® are added to a 60 mm dish and the cells are scraped off using a rubber policeman.

Then split into two 1.5 ml eppendorf tubes and incubated at room temperature for 2 minutes. Tubes are centrifuged at 10,000 x g for 2 minutes to remove the cell debris, and incubated for another 5 minutes at room temperature. Then 200 pi of chloroform is added for each 1 ml of TRIzol® used, mixed by vortexing for

15 seconds, and incubated on ice for 5 minutes. The RNA is extracted into the aqueous phase which is completely separated by centrifugation at 14,000 x g for

5 minutes at 4°C. This aqueous layer is removed and extracted with one volume of phenol:chloroform:isoamyl alcohol (25:24:1) and again with an equal volume of chloroform:isoamyl alcohol (24:1). Sodium acetate, pH 5.2 is added to 0.3 M along with 2.5 volumes of ice cold 100% ethanol. The RNA is precipitated by incubating for 20 minutes at -80°C.

2.15 Isolation of total protein from frog tissues.

One female Xenopus laevis is dissected and tissues are removed and treated as follows. The liver and lung are perfused with 1 x 88 0. The muscle and oviduct are washed with 1 x 880, and the intestine is washed with 1 x 8 8 0 and cut open to expose the inside wall which is then scraped clean to remove any contaminates using a glass cover slip. Afterwards it is again rinsed with 1 x

57 SSC. Oocytes are prepared as described in section 2.14.2. Then all of the tissues are blotted dry and weighed. After weighing they are placed into a petri

dish, chopped into 1 mm cubes, and 1.3 ml of homogenizing buffer (40 mM Tris-

HCI, pH 7.5, 10 mM MgCIa, 7% sucrose, 2 mM DTT, 0.2 mM PMSF, 0.5 pg/ml

leupeptin, 0.7 pg/ml pepstatin A, 2 pg/ml aprotinin, 50 mM NaF, 0 . 2 mM NaVOs)

is added. Tissues are then homogenized with a Wheaton Overhead Stirrer and tapered tissue grinder with Teflon® pestle at a speed of 4000 RPM for two

minutes at 4 °C. The homogenized lysate is centrifuged at 10,000 rpm in a

Sorvall SA 600 fixed angle rotor for 30 minutes at 4 °C, the supernatant is

removed, and protein is quantitated by Bradford assay. However the ovoduct supernatant needs to be sonicated 3 times at a setting of 5 for 30 seconds using

a sonic dismembrator 60 from Fisher Scientific (Pittsburgh, PA), in order to

decrease the viscosity of this sample. Then spun again at 10,000 rpm in a

Sorvall SA 600 fixed angle rotor for 30 minutes at 4 °C. This supematant is also

removed, and quantitated by Bradford assay.

2.16 Isolation and maturation of oocytes.

Oocytes are obtained from the ovaries and separated from the follicle cells by manual dissection as described in Methods in Cell Biology Volume 36 (165).

The isolated oocytes are then incubated ovemight in OR2 medium (5 mM

HEPES, pH 7.8, 82.5 mM NaCI, 2.5 mM KOI, 1 mM CaClg, 1 mM MgClg, 1 mM

Na 2 HP0 4 ) with or without 10 pg/ml of progesterone. The oocytes are then removed from the medium and processes as described in section 2.14.1.

58 2.17 Isolation of total protein from mouse tissues.

One male mouse is dissected and the tissues are removed. The liver and lung are perfused with 1 x SSC. The muscle, Kidney, and spleen are washed with 1 X SSC. While the intestine is washed with 1 x SSC, cut open, scrapped with a glass cover slip, and finally rinsed with 1 x SSC. All the tissues are blotted dry and weighed, then chopped into 1 mm cubes. Then 1.3 ml of homogenization buffer (40 mM Tris-HCI, pH 7.5, 10 mM MgClg, 7% sucrose, 2 mM DTT, 0.2 mM

PMSF, 0.5 pg/ml leupeptin, 0.7 pg/ml pepstatin A, 2 pg/ml aprotinin, 50 mM NaF,

0.2 mM NaVOa) is added for each gram of tissue. Tissues were then homogenized using a Wheaton Overhead Stirrer and tapered tissue grinder with

Teflon® pestle at a speed of 4000 RPM for two minutes at 4 °C. The homogenized lysate is centrifuged at 10,000 rpm in a Sorvall SA 600 fixed angle rotor for 30 minutes at 4 °C, and the supematant is removed and protein is quantitated by Bradford assay.

2.18 Activity assays with frog and mouse tissues.

A 5’pP]labeled transcript corresponding to nt 1690-2002 of the albumin mRNA is prepared as described previously (163). This is incubated with protein extract in a buffer containing 30 mM Tris-HCI, ph 7.5, 1 mM DTT, 2 mM

MgClz and 75 mM KCI, at 25°C using either 5 pg of the frog protein or 0.1 pg of mouse protein, and 1 pi of RNase inhibitor. The reactions are stopped by adding one volume of stop solution (98% formamide (v/v), 0.1% (v/v) bromophenol blue

59 and 0.1 % xylene cyanole) and heating at 95 °C for 3 minutes. The samples are then electrophoresed on a 6 % polyacrylamide/urea gel, and cleavage products are visualized by autoradiography.

2.19 Calf Intestine Alkaline Phosphatase treatment of protein samples.

Ten pg of total protein is diluted to 25 pi with 10 mM Tris-HCI, pH 8.0 and

25 U of CIAP. Thisias incubated for 30 minutes at 37 °G then 5 pi of 5 X SDS-

PAGE loading buffer is added and the sample is heated at 100 °C for 5 minutes and then subjected to western blotting as described in section 2.5. Blots are probed with a polyclonal PMR - 1 antibody (ABa).

2.20 Isolation of total protein from cell culture.

Cells are rinsed in 1 ml of PBS, scraped off the dish, and pelleted by centrifugation in a serofuge. The pellet is flash frozen and then dissolved in 2 ml of homogenizing buffer (40 mM Tris-HCI, pH 7.5, 1 0 mM MgClg, 7% sucrose, 2 mM DTT, 0.2 mM PMSF, 0.5 pg/ml leupeptin, 0.7 pg/ml pepstatin A, 2 pg/ml aprotinin, 50 mM NaF, 0.2 mM NaVOs). Cells are then homogenized by 20 strokes of a 7 ml dounce homogenizer and centrifuged at 10,000 rpm in the SA

600 rotor at 4 °C for 30 minutes. The supematant is removed and concentrated by adding 3 volumes cold acetone followed by incubation at -20 °C for 15 minutes. Samples are then centrifuged at 10,000 rpm for 15 minutes and the pellet is dissolved in sterile water, and quantitated by Bradford assay.

6 0 2.21 Ligation Mediated Polymerase Chain Reaction (LM-PGR).

2.21.1 Primer ligation: DMSO method.

RNA-primer ligation reactions containing DMSO were performed in a 15 pi volume containing 2-10 pg of total RNA and 2 pg of ligation primer M H IIN H 3 P in

50 mM HEPES, pH 8.3, 10 mM MgClg, 5 mM DTT, 2 mM ATP, 50 pg/ml BSA,

25% (v/v) DMSO and 30 units placental ribonuclease inhibitor. The ligation primer MH11NH3P (P-CCAGGTGGATAGTGCTCAATCTCTAGATCG-NH 3 ) was prepared by Operon, and has 5’ phosphate and 3' amine termini, respectively.

Ligation was performed at 4 °C for 16 hr with 15 units of T4 RNA (Life

Technologies). The reaction mixture was extracted once with one volume of phenol:chloroform:isoamyl alcohol (25:24:1) and once with one volume of chloroform:isoamyl alcohol (24:1). The aqueous layer was adjusted to 0.3 M sodium acetate, pH 5.5, and RNA with the ligated primer was precipitated by addition of 2.5 volumes of ice-cold ethanol.

2.21.2 Primer ligation: PEG method.

Between 2 and 10 pg of total RNA was added to a 15 pi reaction containing 50 mM Tris, pH 8 , 10 mM MgClg, 20 mM ATP, 2 mM DTT, 10 pg/ml

BSA, 1 mM hexamine cobalt chloride, 25% (w/v) polyethylene glycol 8000, and

30 units placental ribonuclease inhibitor and 2 pg of ligation primer MHIINH3P.

The primer MHIINH3P (P-CCAGGTGGATAGTGCTGAATGTGTAGATCG-NH3) was prepared by Operon, and has a 5’ phosphate and 3’ amino termini.

Ligations were performed at 4 °G 16 hr with 15 units of T4 RNA ligase (Life

61 Technologies, Inc.)- The reaction mixture was extracted once with one volume of

phenol:chloroform:isoamyl alcohol (25:24:1) and once with one volume of chloroform:isoamyl alcohol (24:1). The aqueous layer was adjusted to 0.3 M sodium acetate, pH 5.5, and RNA with the ligated primer was precipitated by addition of 2.5 volumes of ice-cold ethanol.

2.21.3 RT-PCR.

The entire mixture of RNA ligated to the MH11NH3P primer was

resuspended in 11 pi of DEPC-treated water, followed by addition of 250 ng of the primer MH12 (5’CGAGCTAGAGATTGAGCAC), which is complementary to the first 19 nt of MH11NH3P. The solution was heated to 100 °C for 3 minutes and quickly cooled on ice. To this was added 4 pi of 5X buffer (250 mM Tris-HCI, pH 8.3, 375 mM KCI, 15 mM MgClg), 10 mM DTT and 0.5 mM dNTPs) and HgO to a total volume of 20 pi. The mixture was heated at 42 °C for 2 min followed by the addition of 200 units of Superscript II reverse transcriptase (Life

Technologies, Inc.), and incubated for an additional 50 min. The reaction was terminated by heating at 70 °C for 15 min. Three pi of the above reaction mixture was mixed with 2.5 pi of 10X buffer (100 mM Tris-HCI, pH 8.9, 1 M KCI,

15 mM MgClz, 500 pg/ml BSA, 0.5% Tween 20 (v/v)). To this was added MgClg and dNTP’s to a final concentration each of 3 mM. The wax bead of the Hot

Start™ tube (Life Technologies, Inc.) was melted by heating the tube to 75 °C for about 30 seconds and the mixture was cooled quickly on ice. 11 pi of DEPC water, 2 pi of 5' [^^P] labeled gene specific primer, and 0.5 pi of tTh polymerase

62 from Roche Molecular Biochemicals (Indianapolis, IN) were then added to a final volume of 25 pi. The mixture was heated at 95 °C for 2 min and PCR amplification was performed for 25 cycles at 95 °C for 2 min, 58 °C for 30 sec, and 72 °C for 1 min, followed by extension for 3 min at 72 °C using primers synthesized by Operon Technologies (Alameda ,CA). The gene-specific primers

used were. Set A1 (5’CGCGGTACCTGGATCACCCTGATTTGTC, beginning at position 40 of albumin mRNA), Set G1 (5'TCCTTGTGAAGCTGATTA, beginning at position 1690 of albumin mRNA), and MH28 (5’AAGAGGCGAACACACAACG, beginning at position 1669 of c-myc mRNA). The reaction mixtures were then extracted as above with phenol:chloroform:isoamyl alcohol, and amplified products were recovered by ethanol precipitation. The recovered pellet was dissolved in 3 pi of DEPC H 2 O and an equal volume of formamide loading dye

(80 % formamide, 1 mM EDTA pH 8, 0.1 % bromophenol blue, and 0.1 % xylene cyanol). This was heated for 5 min at 95 °C and loaded into a single lane of a denaturing 6% polyacrylamide/urea gel.

2.21.4 Product recovery, reamplification and sequencing.

Products from the above reaction were identified by autoradiography of the dried polyacrylamide gel. The bands of interest were extracted from the dried gel following a modified protocol from Jo et. al. (166). The film was aligned with the gel and an 18 gauge needle was used to punch holes at the 4 comers of each band. A fresh razorblade was then used to cut out each desired band, and the excised gel fragments were soaked in 200 pi of distilled HgO for 10 minutes.

63 The tube was then boiled for 15 minutes with parafiim to hold the lid closed. This

was then centrifuged at 10,000 x g for 2 min and the supematant was removed.

DNA was recovered by addition of 1/10^ volume of 3 M sodium acetate, pH 5.5,

50 pg glycogen, and 900 pi of 100% ethanol, followed by chilling at -8 0 °C for 30

min. The DNA pellet was recovered by centrifugation at 14,000 x g, 4 °C for 15

minutes, washed with cold 85% ethanol and dissolved in 12 pi of distilled H 2 O.

Four pi of the extracted DNA was added to a 40 pi reaction containing 4 pi of a 10X buffer consisting of 100 mM Tris-HCI, pH 8.9, 1M KCI, 15 mM MgCIa,

500 pg/ml BSA, 0.5% Tween 20 (v/v). The reaction mixture was adjusted to 1.5 mM MgClg, followed by the addition of 0.4 mM dNTP’s, 1 ng of primer MH12, 1 ng of the gene specific primer, and 2.5 U of tTh polymerase (Roche Molecular

Biochemicals, Inc.). PCR amplification was performed as described in section

2.17.3, following which 8 pi was treated with 10 units of exonuclease I and 2 units of shrimp alkaline phosphatase for 20 minutes at 37 °C to remove unincorporated primers. This reaction was terminated by heating for 15 minutes at 80 °C, followed by addition of 20 ng of the original gene-specific primer. This was annealed to the template by heating at 100 °C for 3 minutes and then cooling on ice for 5 minutes, and sequenced using T7 DNA polymerase (USB T7

Sequenase Kit). The products were denatured and electrophoresed on a 6% polyacrylamide/urea gel as described in section 2.17.3.

64 2.22 In vitro activity assay.

A 5’[^^P]labeled transcript corresponding to nt 1690-2002 of the albumin

mRNA is prepared as described previously (163). This transcript is incubated

with protein extract in a buffer containing 30 mM Tris-HCI, ph 7.5, 1 mM DTT, 2

mM MgCIa and 75 mM KCI, at 23“C using either 10 pg of unfractionated

polysome extract from estrogen-treated frogs (Fig 3.17A), or 20 units of PMR-1

purified as described previously (129) (Fig 3.17B). One unit of PMR-1 is the amount needed to completely cleave 7 fmol of albumin substrate transcript in 30

min at 23 °C. The reactions are stopped by adding one volume of stop solution

(98% formamide (v/v), 0.1% (v/v) bromophenol blue and 0.1% xylene cyanole) and heating at 95 °C for 3 minutes. The samples are then electrophoresed on a

6% polyacrylamide/urea gel, and cleavage products are visualized by autoradiography.

2.23 RNase T1 digestion.

A 5’ [^^PJIabeled transcript corresponding to nt 1690-2002 of albumin mRNA was added to 10 pi of hybridization III buffer from the Ambion RNase

Protection Assay III kit (Ambion, Austin, TX). This was heated for 5 minutes at

50 °C, and cooled slowly to room temperature. To this was added 150 pi of

RNase digestion buffer III containing 5 units of RNase T1. Samples were incubated for 5-25 minutes at 37°C and the reaction was stopped by addition of

225 pi of RNase inactivation/ precipitation III solution. Digested RNA was recovered by addition of 150 pi of ethanol, and 10 pg of yeast tRNA, followed by

65 precipitation at -20 °C . The pellet was dissolved in 6 pi of gel loading buffer II and products were separated on a 6% denaturing polyacrylamide/urea gel and visualized by autoradiography. The positions of single stranded G residues identified by RNase T1 digestion were used to model the secondary structure of this portion of albumin mRNA using the MFOLD server (167).

2.24 Isolation of cytoplasmic protein from hepatocytes.

It is important to keep everything cold so after scraping the cells all steps are performed in the cold room. The medium is removed from the primary hepatocytes which are then washed with 2 ml of PBS + glucose. Cells are scraped off the plate with a rubber policeman and pipetted into a 3 ml cell culture tube, centrifuged in a serofuge to remove the matragel. The cell pellet is then dissolved in 1 ml of PBS + glucose and centrifuge at ^10,000g for 30 seconds at

4 °G. Discard the supernatant and add 200 pi of extraction buffer (10 mM Tris-CI, pH 8.6, 0.14 M NaCI, 1.5 mM MgCI2, 0.5% Nonidet P-40 (NP-40), 1 mM dithiothreitol (DTT), 25 pl/ml sigma protease and 10 pl/ml sigma phosphatase inhibitors) vortex for 15 seconds and incubate on ice for 5 minutes. Then centrifuge the cells at 12,000g for 90 seconds and keep the supematant which is your total protein.

2.25 Xenopus primary hepatocyte cell culture.

Male Xenopus livers are removed as described in section 2.1 without the estrogen treatment, chopped into 1 mm cubes and disaggregated with rocking for

6 6 several hours at room temperature in Km buffer (10 mM HEPES, pH 7.3, 180 mM NaCI, 2 mM KCI, 4 mM sodium pyruvate, 9 mM anhydrous D-glucose, 0.5%

BSA) supplemented with 400 g/ml collagenase, 200 unites/ml penicillin and 200 g/ml streptomycin. Cells are recovered by gentile agitation with a 25 ml pipette and removed from the colagenase solution by brief centrifugation at 50 x g. The cells are then fractionated first using a 25% and 50% Percoll step gradient centrifuged for 10 minutes at 50 x g in KM buffer. Hepatocytes and melanocytes sediment to the bottom of the tube. Then using a 60% and 90% Percoll step gradient centrifuged for 10 minutes at 300 x g in KM buffer the hepatocytes sediment at the interface between the 60% and 90% Percoll. The cells are washed in Km buffer supplemented with penicillin and streptomycin (as above) three times and centrifuged at 50 x g to pellet the cells, to remove the Percoll.

The final cell pellet is dissolved in complete 0.65 X phenol red free Coon's modified Ham’s F I2 medium containing 20 mM HEPES, 2.6 g/L NaHCOa

(adjusted to pH 7.3 with NaOH), 100 units/ml gentamicin, 100 nM dexamethasone, 10 g/ml insulin, and 10 nM L-triiodothyronine. Typically 1.7 million cells are plated onto 60 mm tissue culture dishes coated with 2 ml of a chilled 1:10 dilution of phenol red free Matrigel™ (Collaborative Research,

Bedford, MA). The cells are then placed in a humid environment at 25 °C and half the media is changed every 24 hours. Cells treated with 1 pM moxestrol, a poorly metabolized derivative of estrogen, have half the media replaced with media containing 2 pM moxestrol every 12 hours.

67 2.26 Transfection of hepatocytes with antisense Morpholino oligonucleotides.

Morpholino ollgos are used in concentrations from 0.025 pM to 1 pM and transfected into hepatocyte cells one day after being plated using the Effectene™

Reagent from Qiagen (Valencia, CA). Since the Morpholino backbone is uncharged it must be annealed to a complementary DNA oligonucleotide for the lipid reagent to be bind the DNA and transfect it into the cells. This is done by incubating equal amounts of the Morpholino oligonucleotide, PMR-1 antisense

(AS-PMR (TGCAGGAGCCATTTTCTGTAGCACA)), or a Fluorescent labeled

Standard Control (Std-control (GCTCTTACCTCAGTTACAAI I lATA)) and a complementary DNA oligo, MH33 (G AAAAT G G CTCCTGC AAAAAAAAAAA), or

MH32 (TAACT G AG GTAAG AG G AAAAAAAAAA) respectively. It is important to have the Poly(A) tail on the complementary oligonucleotide. This is then heated to 100°C for 3 minutes and quickly cooled on ice. This dsDNA is transfected into the cells following the instructions from the Effectene™ Kit as follows. The DNA is diluted to 150 pi with buffer EC, and incubated with 8 pi of Enhancer for 5 minutes at 25 °C. Then 25 pi of Effectene™ Transfection reagent is added, and incubated at room temperature for 10 minutes. While this is incubating the medium is aspirated from the 60 mM dishes and 4 ml of fresh medium is added.

Afterwards 1 ml of growth medium is added to the reaction tube, mixed briefly and added drop-wise to the cells. Gently swirl the dish to distribute evenly. Due to the negative effects of the Effectene™ Reagent on cell viability over long

6 8 periods of time, after 2 hours the cells are washed once with PBS + glucose and fresh medium in added. RNA is isolated 48 hours later.

2.27 Isolation of Xenopus hepatocyte cytoplasmic/nuclear RNA.

Medium is removed from the plates and 1.5 ml of PBS + glucose with 10 U of RNase inhibitor is added. A rubber policeman is used to scrape the cells off the plate and pipetted into a 3 ml cell culture tube on ice. Centrifuge the cells for

30 seconds in the serofuge to remove the matragel. Wash the cells with 1 ml of

PBS + glucose with 10 U RNase inhibitor, again centrifuge in the serofuge for 30 seconds. Resuspend the pellet in 200 pi of RNA extraction buffer (10 mM Tris-

HCI, pH 8.6, 0.14 M NaCI, 1.5 mM MgCI2, 0.5% Nonidet P-40 (NP-40), 1 mM

DTT, 1 U/100 pi RNase inhibitor) and vortex. Incubate on ice for 5 minutes and centrifuge for 1.5 minutes at 10,000 x g. Transfer the supernatant (cytoplasmic portion) to a new tube with 1 ml TRIzol® reagent. Also add 1 ml TRIzol® reagent to the pellet and resuspend. Incubate both at 25 °C for 5 minutes then add 200 pi of chloroform for each ml of TRIzol®. Incubate another 5 minutes and centrifuge 15 minutes at 12,000 x g. Remove the aqueous phase and add 0.5 ml of isopropanol per 1 ml of TRIzol® used and Sodium acetate, pH 5.2 is added to

0.3 M. Precipitate the RNA by incubating at -20 °C for a minimum of 30 minutes.

2.27.1 Northern blot.

2 pg of total RNA is diluted into a 12 pi volume of sterile water. Add 3 pi of deionized glyoxal and 2 pi of 0.1 M phosphate buffer. Heat at 50 °C for no

69 more than one hour. Cool the RNA samples to room temperature and add 4 pi of 5X loading dye (50% glycerol, 0.01 M sodium phosphate, pH 7.0, 0.4% each bromophenol blue and xylene cyanole). The gel boat and comb are washed in 1

N NaOH for 20 minutes, and a 1% (w/v) agarose gel is used. Electrophorese at

100 V for approximately 3 hours, until the bromophenol blue is about 1 cm from the bottom of the gel. Once both dyes have entered the gel start the stir bar at low speed. Remove the gel and blot onto Nytran membrane using the Turbo

Blotter (Scheicher and Schuell; Keene, NH) as described by the manufacturer with 10x SSC. After blotting rinse the blot with 6x SSC and allow to air dry for 30 minutes. Crosslink the RNA to the Nytran membrane by exposing to a 254 nm

UV-lamp (Stratalinker® 1800, at 120,000 pJ/cm^) for 45 seconds.

2.27.2 Hybridization of Northern blot.

The blot is prehybridized with 7 ml of Sigma Perfecthyb plus solution for at least 30 minutes at 68°C. Ten million cpm of each probe are then added directly to the perfecthyb plus solution and incubated at 68 °C overnight. The next morning the blot is washed twice with low stringency buffer (1% SDS, 2 x SSC) for 10 minutes at 25 °C, twice with high stringency buffer (1% SDS, 0.5 x SSC) for 20 minutes at 68 °C, and once with ultra high stringency buffer (1% SDS, 0.1

X SSC) for 2 hours at 68 °C.

70 2.27.3 Labeling probe.

Probes for the northern blots are made using the Random Primer DNA labeling kit from Life Technologies. 25 ng of DNA dissolved in 20 pi of distilled water is denatured by heating for 5 min in a boiling water bath, and immediately cooled on ice. Keeping the DNA on ice 2 pi dATP solution (0.5 mM dATP in 3 mM Tris-HCI (pH 7.0), 0.2 mM NagEDTA), 2 pi dGTP solution (0.5 mM dGTP in 3 mM Tris-HCI (pH 7.0), 0.2 mM NagEDTA), 2 pi dTTP solution (0.5 mM dTTP in 3 mM Tris-HCI (pH 7.0), 0.2 mM NaaEDTA), 15 pi Random Primers Buffer Mixture

(0.67 M HEPES, 0.17 M Tris-HCI, 17 mM MgCI2, 33 mM 2-mercaptoethanol,

1.33 mg/ml BSA, 18 OD260 units/ml oligodeoxyribonucleotide primers

(hexamers), pH 6.8), 5 pi (approximately 50 pCi) [alpha-32P]dCTP, 3000

Ci/mmol, and distilled water is added to a total volume of 49 pi. Add 1 pi Klenow

Fragment (3 units), mix gently but thoroughly, and centrifuge briefly. Incubate at

25 °C for 1 h. Incubation times longer than 1 h may give higher specific activity.

Add 5 pi of the stop buffer (0.2 M Na2EDTA, pH 7.5).

The labeled probe is then cleaned up using a G25 spin column from 5- prime, 3-prime as follows. First the column resin needs to be resuspend and allowed to settle by draining off the buffer. Place the column with a 1.5 ml microfuge tube into a 15 ml conical tube and centrifuge at 1,100 x g for 1 minute in the hermie Z320 centrifuge with a swinging bucket rotor. Empty the collection tube and centrifuge at 1,100 x g for another minute. Place the column into another collection tube and carefully apply the labeling reaction directly to the top

71 of the resin. Centrifuge at 1,100 x g for 4 minutes, the probe will be in the collection tube and greater than 99% of the unincorporated nucleotides should be trapped in the resin. The amount of incorporated counts can be determined by spotting 1 pi of the flow through on a glass fiber filter, letting it air dry and washing with 50 ml of ice cold 10% (w/v) TCA containing 1% (w/v) of sodium pyrophosphate. Then a scintillation counter can be used to determine the counts on the membrane, or incorporated counts.

72 CHAPTER 3

RESULTS

3.1 Biochemical comparison of hMPO and PMR-1.

3.1.1 The absence of heme distinguishes PMR-1 from hMPO.

When PMR-1 was cloned it appeared to be a member of the peroxidase gene family by its 57% sequence identity to hMPO. The aligned sequences of both proteins are shown in Figure 3.1 A. With the exception of the very N and C- termini there is sufficient sequence identity that one might assume the protein we cloned was Xenopus myeloperoxidase. We therefore sought to determine whether this was the case, using several unique properties of hMPO as criteria for comparison of the two proteins. hMPO is a heme-containing protein in which iron protoporphyrin IX is covalently bound at 3 sites to the apoprotein (159).

These sites (E408, M409, D260) are indicated in larger bold type in Figure 3.1 A, and the structure of the bound heme complex is shown in Figure 3.2A. Whereas

E408 and D260 are conserved between hMPO and PMR-1, M409 is replaced in the latter by glutamine. The same substitution is seen in other peroxidases such as lactoperoxidase. This difference provided a unique opportunity to compare these proteins, as the heme attachment at M409 consists of a novel sulfonium ion bond that can be activated upon heating to produce cleavage of the peptide

73 Figure 3.1 Sequence and structural features of PMR-1. A. The sequence of PMR-1 (GenBankAccenssion Number U68724) is aligned with human myeloperoxidase (hMPO), with identical residues identified with grey boxes, and the tryptic peptides of purified PMR-1 indicated with open boxes. The black underline denotes the hMPO signal sequence, the open underline the hMPO propeptide, the lightly lined underline the hMPO light chain, and the darkly lined underline the hMPO heavy chain. The sites of covalent heme attachment to hMPO are shown with an asterisk (•) ant the sites that coordinate the heme iron are shown with a black dot (•). An open arrow denotes the proposed proteolytic processing site that generates mature PMR-1. B. Ten micrograms of total liver protein obtained 24 h after injection of estradiol were analyzed by Westem blot using a polyclonal antibody to purified PMR-1 (88). Markers indicate the presumptive 80-kDa precursor, mature PMR-1, and a 40-kDa breakdown product commonly observed on SDS-PAGE (88).

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501 1 600 B a a oasLzgarzT auD G ntna wmtaswam ttiiaim rr— a s z D n u n . m m k u o m s agLW Dua uwLnou QfpaOÊnæ wxanaomm momvfLSRv rraj—m jt oazpaiHBL magMiTO iPSPATMat i______

6 0 1 , 7 0 0 MBRfCBLSA tSMVKBUUMT ZMNnOMOO ZK LW IB S D I«V m «8 IVM RZO K^ ZSCUawgVK RAIBM g iraüPBVWWBQl BASZPVnA A— Af CQUQ fBTVOQZOr? tiOBKLAeSL M iflZO iB H DZMWMET IKBMKVqW, EAC lU ggn t 6T.»qpa M I BttOVnMQg aQALAQZSLF

7 0 1 , 7 4 5 ayzoB pgg BVPRiioiirO ia gp»pvAC a r i a t i p u o w k v a . # residues coordinating heme iron RZZCBWIGZT TvaxmziMB MAlAAtiBnecSTLlAXiiLAS WMAA ^ covalent heme attachment site

B

hepatocyte extract

SOIcDa 60kOa — 40kDa

Figure 3.1 Sequence and structural features of PMR-1

75 backbone in the same manner that CNBr cleaves at methionine residues (164).

This will result in the production of a 20 and 40 kDa fragment from the hMPO heavy chain. Such cleavage of PMR-1 might explain the variable presence of a

40 kDa breakdown product in addition to the 62/64 kDa doublet seen on SDS-

PAGE of PMR-1 Figure 3.1 B.

The upper westem blot in Figure 3.2B shows the result of autolytic cleavage experiments performed on hMPO. In lane 4 purified hMPO was incubated with SDS sample buffer containing |3-mercaptoethanol to reduce the disulfide bond linking the light and heavy chains, but was not heated prior to application to the gel. Seen here are the -60 kDa glycosylated heavy chain, and the 15 kDa light chain. Lanes 1 -3 show the effects of various combinations of heating hMPO in the presence or absence of the reducing agent. In each case, heating at 95°C caused autolytic cleavage of the heavy chain of the protein to yield 40 and 20 kDa fragments. No fragmentation was seen for the light chain.

The lower western blot show the effects of the same treatment regimens on

PMR-1 present in a polysome extract that was processed through the first step of purification on a QAE column. In keeping with the substitution of glutamine for methionine at 409, no autolytic cleavage was seen for PMR-1.

As mentioned previously lactoperoxidase also has covalently-bound heme, but it lacks the methionine at position 409 and therefore does not have the sulfonium ion linkage between the heme and the protein (168). This results in

76 Figure 3.2 PMR-1 does not undergo autolytic cleavage upon heating. A. Structure of the m heme of myeloperoxidase. B. 10% SDS-PAGE gel of hMPO that has been Incubated with [3-mercaptoethanol and /or heated. Treatment with (3-mercaptoethanol leads to separation of the heavy (60 kDa) and light (15 kDa) while heating leads to autolytic cleavage of the heavy chain (40 and 20 kDa) between methionine 409 and proline 410 resulting from the formation of a methionyl sulfonium intermediate with methionine 409 and the porphyrin prosthetic group. The bottom gel shows PMR-1 that has been treated the same as the hMPO samples.

77 Met409

CH3

CH. GIU408

Fe.

Asp260

COOH COOH

liJ

0 0 C O . s B

40 kDa -

hMPO 20 kDa -

15 kDa

PMR-1

1 2 3 4

Figure 3.2 PMR-1 does not undergo autolytic cievage upon heating.

78 Figure 3.3 Wavelength scan of PMR-1 and hMPO. Either hMPO (--) or PMR (—) were diluted In 50 pi of a 40 mM TrIs-CI, pH 7.5, 2 mM DTT buffer to 0.1 pg/ml or 0.02 pg/ml respectively. These were then scanned from 200 nm to 600 nm using a Beckman DU 640 spectrophotometer.

79 0.150

8 g -Eo (/) hMPO .Q (ü

PMR-1 0.00 200 300 400 500 600 wavelength (nm)

Figure 3.3 Wavelenth scans demonstrate that unlike hiMPO, PMR-1 does not have a heme group.

8 0 different spectral properties; lactoperoxidase has a So ret band at 413 nm whereas that of hMPO is red-shifted to 430 nm. To determine whether PMR-1 contains heme the UV-visible spectrum of PMR-1, purified from liver polysomes of estrogen treated frogs as described previously (129), was compared with that of hMPO. hMPO displays the characteristic absorbance of the So ret band at 430 nm and a secondary absorbance at 570 that has been noted previously (168). In contrast, PMR-1 shows no absorbance in the visible spectrum. If PMR-1 had heme bound in the same manner as lactoperoxidase, one would have seen absorbance at 500 and 550 nm in addition to the 413 lactoperoxidase Soret band. These results indicate that PMR-1 is distinct from hMPO and other members of the peroxidase gene family by the absence of covalently bound heme.

3.1.2 hMPO is glycosylated whereas PMR-1 Is not.

As noted above, hMPO is a glycoprotein. Figure 3.4 lists all of the potential sites for N-linked glycosylation of both hMPO and PMR-1. Nauseef determined hMPO has 5 N-linked high mannose chains (158).

Since the glycosylation site at hMPO position 139 is not present in the fully- processed protein, the 5 sites of N-linked glycosylation must correspond to the remaining 5 shown in Figure 3.4. A scan of PMR-1 indicates the presence of 6 putative N-linked glycosylation sites, 3 of which (326, 362 and 453) are homologous to sites in hMPO. Two approaches were used to evaluate whether there is N-linked glycosylation of PMR-1 ; cleavage of the sugar backbone with

81 endoglycosidase H (Endo H) and affinity chromatography on concanavalin A- sepharose (Con A sepharose). Endo H is an endoglycosidase that specifically cleaves between the two N-acetylglucosamines present in the core of high mannose oligosaccharides. Complex oligosaccharides are resistant to Endo H cleavage. In the experiment shown in Figure 3.5 hMPO was incubated without and with Endo H, then processed for SDS-PAGE including heating at 95°C which produces the autolytic cleavage of the heavy chain as noted above. The resultant product was then separated on a 10% gel, and analyzed by Westem blot using a polyclonal antibody to hMPO. Each of the species noted in Figure

3.2 are seen in the control lane. Upon Endo H digestion the bands corresponding to the 60 kDa heavy chain (filled arrow) and the 40and 20 kDa autolytic cleavage products show increased mobility indicative of removal of covalently bound carbohydrate. The difference is most striking for the 20 kDa fragment, which, after Endo H digestion, migrates with the same mobility as the non-glycosylated 15 kDa hMPO light chain. In the experiment on the right side of the figure, 10 and 20 micrograms of polysome extract containing PMR-1 were treated with Endo H and separated on an SDS-PAGE gel and detected using a polyclonal antibody raised against PMR-1 (ABa). No shift in mobility was evident with Endo H treatment, indication that PMR-1 lacks the high mannose oligosaccharides that are present on hMPO.

These data do not rule out the possibility that PMR-1 bears Endo H resistant complex oligosaccharides. Concanavalin A binds oligosaccharides

82 Figure 3.4 N-iinked glycosylation sites on hMPO and PMR-1 Potential N-lInked glycosylation sites on PMR-1 were identified using MOTIFS in the Wisconsin GOG program. N-linked glycosylation sites on hMPO were obtained from the Brookhaven Protein Database (ImhI) for the protein structure at 2.25 Â resolution. The position of the first amino acid of each region is denoted, with the putative N-linked glycosylation signal (N-X-S/T) indicated in bold.

83 Position hMPO sequence PMR-1 sequence Position

323 ACPGSNITIRNQIN DPRISNQSDCIPLF 277

355 ARNLNMSNQLGLL AVKLRNNTNQLGLM 326

391 PGLLTNRSARIPCF FCVLTNRSSGIPCF 362

483 TYRSYNDSVDPRIA AYRSYNESVDPRVS 453

Figure 3.4 N-linked glycosylation sites on hMPO and PMR-1

84 Figure 3.5 Endoglycosidase H digestion shows no high mannose oligosaccharides on PMR-1. Either 1 |jg of hMPO, 10 of partially purified PMR-1 extract, or 20 pg of partially purified PMR-1 extract were incubated at 37 0 for 1 hour with or without 750 units of endoglycosidase H. The extracts were then electrophoresed on a 10% SDS-PAGE gel. The size of the markers are shown on the side of each gel.

85 polysome extract hMPO 10 20 |Lig — 97

H — 66 — 45 kOa kDa

31 — 31 21.5 — < = ■ 21.5

L > - + endo H

Figure 3.5 Endoglycosidase H digestion shows no high mannose oligosaccharide on PMR-1.

8 6 Figure 3.6 Concanavalin A-Sepharose binding shows no oligosaccharides on PMR-1. 10 ijg of partially purified PMR-1 extract was incubated with concanavalin A bound to sepharose beads for 20 minutes at 4°C and the supernatant was removed. The beads were washed with buffer three times and then washed with buffer and 500 mM methyl-a-D-mannopyroside to elute anything that was bound to the concanavalin A. TCA precipitated samples were then electrophoresed on a 10% SDS-PAGE gel which was transferred to an Imobilon P membrane and western blotted using monoclonal hMPO or polyclonal PMR-1 antibodies. Sizes of molecular weight markers are shown on the left side of the gel.

87 wash eluate hMPO 80 51 ■ H 34 kDa 27—

16.6 —

PMR-1 80 — 51 — 34 — kDa 27—

16.6 —

1 2 3 4 5 6 7 8

Figure 3.6 Concanavalin A-Sepharose binding shows no oligosaccharides on PMR-1.

8 8 bearing either a—D-mannosyl or a-D-glucosyl residues present in both high mannose and complex oligosaccharides. We therefore used binding to Con A sepharose to determine whether PMR-1 differs from hMPO by having complex rather than high mannose oligosaccharides, or whether it lacks glycosylation completely. In the experiment in Figure 3.5 hMPO or a partially purified preparation of PMR-1 were incubated batchwise with 40 pi of Con A sepharose.

After washing to remove unbound material, bound glycoprotein was eluted with a-methyl-D-mannosylpyranoside. The fractions obtained from this treatment were analyzed by westem blot using either a monoclonal antibody to the hMPO heavy chain, or an epitope-specific antibody to PMR-1. The data in Figure 3.6 show that a considerable amount of input hMPO remained unbound in this procedure. However, specific binding of this glycoprotein was evident by elution with a-methyl-D-mannosylpyranoside (lanes 6-8). In contrast, none of the input

PMR-1 bound specifically to Con A sepharose (Fig 3.6 bottom). These results were confirmed by an activity assay using an albumin substrate transcript that showed none of the input PMR-1 bound to the resin (data not shown). We conclude that PMR-1 also differs from hMPO in that it lacks glycosylation of the processed product.

3.1.3 PMR-1 and hMPO have distinct enzymatic activities.

The high degree of structural similarity between hMPO and PMR-1 raised the possibility that the former might possess ribonuclease activity in addition to its peroxidase activity, or that PMR-1 might possess peroxidase activity in addition

89 Figure 3.7 Comparison of the enzymatic activities of hMPO and PMR-1. A. 10 |jg of a partially purified PMR-1 extract (lanelO), or pure hMPO 1,2, or 3 pg with (lanes 7-9) or without 1 mM Mg^^ (lanes 3-5) were incubated for 30 min at 22°C in the presence of uniformly labeled albumin transcript. Lane 1 is labeled markers and lanes 2 and 6 show the input transcripts incubated without added protein. RNA degradation was assessed by electrophoresis of the resulting mixtures on a 6 % urea/acrylamide gel. The characteristic doublet cleavage product of PMR-1 with the albumin transcript is shown by the arrow. B. Duplicate 100 ng of hMPO or 100 ng of PMR-1 purified from liver polysomes were assayed for peroxidase activity using ARTS substrate. The data are shown as O D 4 0 5 generated per microgram of protein.

90 + 1 mM Mg+2

hMPO hMPO PMR-1

123456789 10

B hMPO 20

E 15 2 O) i E 10 § Û O 5 — PMR-1 0

Figure 3.7 Comparison of the enzymatic activities of hMPO and PMR-1

91 to its ribonuclease activity. In the experiment in Figure 3.7A, lanes 2-4, a

[^^P]labeled albumin substrate transcript was incubated with 1, 2, or 3 pg of purified hMPO under the conditions determined previously to assay PMR-1 ribonuclease activity (129). The transcript remained undegraded under these conditions, whereas the same substrate was degraded by PMR-1, resulting in the production of the characteristic doublet cleavage product (lane 10). We noted previously that PMR-1 activity was enhanced by the addition of 1 mM MgClgto the reaction buffer. The addition of divalent cation had no effect on ribonuclease activity of hMPO (lanes 7-9). We conclude that, unlike PMR-1, hMPO lacks ribonuclease activity.

Next we wanted to see if PMR-1 possessed a peroxidase activity like that of hMPO. This was done using the chromogenic substrate ABTS (Fig 3.7B).

Duplicate 100 ng samples of hMPO generated approximately 20 OD 4 0 5 units per pg protein. However PMR-1 produced no such product, demonstrating that

PMR-1 does not have any peroxidase activity.

3.2 Expression of PMR-1 In other tissues and organisms.

3.2.1 PMR-1 antibody cross reacts with a protein In Xenopus and murine tissues.

It is generally believed that there are a small number of endonucleases that are involved in the regulated destabilization of a large number of transcripts in a tissue and environment specific means. The experiment in Figure 3.8A

92 shows that many tissues express a protein that cross reacts with a polyclonal antibody raised against PMR-1 .(ABg) (129). The liver extract demonstrates the characteristic 62/64 kDa doublet of PMR-1 (129). Similar signals are seen in the lung, muscle, intestine, and oocytes. However, the intensity of the signal varies a great deal between these tissues , which suggests that the amount of PMR-1 expressed in each tissue may vary. The only tissue that does not have a protein that cross reacted with ABa was the ovoduct. However, when the membrane was coomosie stained, the ovoduct sample seemed to have about half as much protein as the others (data not shown). This is believed to be due to the viscosity of the ovoduct sample (see materials and methods), which could account for macromolecules that prevented protein from entering the gel. This data suggest that PMR-1 is expressed in most tissues in Xenopus laevis.

Figure 3.88 demonstrates that like Xenopus most murine tissues also express a protein that cross reacts with the polyclonal antibody ABa. Unlike

Xenopus tissue, no doublet was seen in most murine tissues. Also, the protein seems to have a slightly decreased mobility by SDS-PAGE. The same protein also cross reacted with a glycine purified antibody raised against a peptide sequence of PMR-1, 1277 (156). Kirsten Bremer has demonstrated the presence of a protein of similar size in MEL (murine etythroleukemia), MCF-7

(human breast cancer), and HepG2 (human liver) cell lines that cross reacts with

ABa and 1277 (Bremer and Schoenberg personal communication). These data suggest that many mammalian tissues express PMR-1. However, in all of these

93 Figure 3.8 Western blot of frog and mouse tissue shows PMR<1 is present in many tissues. 25 pg of protein isolated from each tissue was separated on a 10% SDS-PAGE gel and visualized by Westem blot using a polyclonal PMR-1 antibody ABa. Size markers are shown on the right. B. 25 pg of protein isolated from each murine tissue (lanes 2-7) or 10 pg of partially purified PMR-1 (lane 1) was separated on a 10% SDS-PAGE gel and visualized by Westem blot using a polyclonal PMR -1 antibody ABa. Size markers are shown at the right

94 V Y 66 1-57 1 2 3 4 5 6

B

1 2 3 4 5 6 7

Figure 3.8 Western bloto f Xenopus and murine tissues show PMR-1 is present in many tissues.

95 cases, the mammalian protein has a decreased mobility by SDS-PAGE when compared to Xenopus PMR-I. This decreased mobility could be explained by altered post translational modification of the protein such as increased phosphorylation, or proteolytic cleavage of the 80 kDa precursor protein at a different site. This could lead to the production of a slightly larger protein compared to Xenopus PMR-1.

3.2.2 Unlike murine tissues,Xenopus tissues all produced the major doublet cleavage of PMR-1.

The experiment in Figure 3.9 shows the standard in vitro activity assay in which PMR-1 has been demonstrated to produce a characteristic doublet cleavage product (169). This experiment was performed with the same Xenopus protein extracts used in Figure 3.8. The characteristic doublet cleavage produced by purified PMR-1 is marked by the arrows on the left. This characteristic cleavage was seen in all of the tissues. This data suggests that

PMR-1 is expressed in all Xenopus tissues. The disappearance of the full length transcript in the oviduct tissue could be due to the presence of a highly active nuclease. Although, it is also possible that the viscous protein sample caused most of the transcript to be retained in the well of the gel.

When the same activity assays were performed with 5 pg of the mouse protein samples, the transcript was completely degraded (data not shown). The reaction was repeated using 1 0 0 ng of total protein, and the time of incubation

96 Figure 3.9 Activity assay of frog tissue proteins demonstrates the presence of active PMR-1 in all tissues. Five pg of protein isolated from each tissue (ianes3-8), or 1 pg of partially purified PMR-1 (lane 2), was incubated with uniformly [32P]labeled 160 transcript for 30 minutes at 25°C. Lane 1 is the transcript incubated without any protein being added. Products were separated on a 6 % polyacrylamide/urea gel. The characteristic doublet produced by PMR-1 is labeled by the arrows at the left.

97 Doublet

1 2 3 4 5 6 7 8

Figure 3.9 Activity assayo f Xenopus tissue proteins demonstrates the presence of active PMR-1 in aii tissues.

98 Figure 3.10 Activity assay of murine tissue proteins suggests that PMR-1 is expressed in mammals. 100 ng of protein Isolated from each tissue (lanes 3-8) or 1 pg of partially purified PMR-1 (lane 2), was incubated with uniformly [32P]labeled 160 transcript for 5 or 30 minutes at 25°C respectively. Lane 1 Is transcript Incubated without any protein being added. Products were separated an a 6 % polyacrylamlde/urea gel and the characteristic doublet produced by PMR-1 Is labeled with the open arrow. The secondary cleavages of PMR-1 that are also seen In the murine tissues are labeled with the closed arrows.

99 Doublet

1 2 3 4 5 6 7 8

Figure 3.10 Activity assay on murine tissue proteins suggests that PMR-1 is expressed in mammais.

1 0 0 was decreased from 30 minutes to 5 minutes (Fig 3.10). Despite these drastic changes, the transcript incubated with protein from the liver, intestine, and kidney was almost totally degraded. None of the samples produced the characteristic doublet cleavage of PMR-1, but they did produce some minor cleavages similar to PMR-1. The same pattern was also seen when PMR 60, which also has a decreased mobility by SDS-PAGE, was expressed in baculovirus. These data suggest that PMR-1 is expressed in murine tissues but potentially is proteolytically processed to produce a larger protein that has a modified specificity compared to Xenopus PMR-1.

3.3 Estrogen and development do not change the effect of CIAP treatment on PMR-1.

PMR-1 has previously been demonstrated to be activated by estrogen in an ER-dependent, but transcription independent manner (153). Estrogen has been demonstrated to activate several kinases such as: p38 MAP kinase

(170,171), JNK (172), Src/Ras/MEK/Erk (173,170), and PKA (174) by non- genomic methods. A similar pathway could account for the rapid induction of

PMR-1 in Xenopus liver. Purified PMR-1 separates into 3 spots on 2D gels.

These could result from phosphorylation or the presence of multiple isoforms.

The latter are commonly seen due to the duplicated Xenopus genome.

Data in the lab using a column purified TAP tag PMR-1, and the anti phospho-tyrosine antibody RC-20, suggest that PMR - 1 is not tyrosine

101 phosphorylated (data not shown). Therefore this work focuses on the possibility of serine/threonine phosphorylation by treating the protein samples with calf intestinal alkaline phosphatase (CIAP), which dephosphorylates all phospho- serine/phospho-threonine residues. This increases the mobility of the protein by

SDS-PAGE. The experiment in Figure 3.11 A shows the effect of CIAP on protein isolated from frog liver after 0, 9, or 24 hours of estrogen treatment. Although

PMR-1 activity peaks 9 hours after estrogen treatment, all three time points showed an equivalent increase in mobility upon CIAP treatment, suggesting that there is no difference in the phosphorylation of PMR-1 with estrogen stimulation.

This was corroborated by Westem blots of 2D gels (Tang and Schoenberg personal communication). Figure 3.11B shows that PMR 60 expressed in baculovirus also has a similar increased mobility, suggesting that it is phosphorylated in a manner similar to PMR-1. Figure 3.11C demonstrates that if

CIAP is heat denatured before incubation with the protein, or an equal molar amount of BSA is added instead, there is no increase in the mobility of PMR-1.

This suggests that the increase in mobility is due to an enzymatic effect of CIAP, as apposed to a physical effect of adding a large amount of protein.

Work done in the Steitz lab has demonstrated that Xenopus oocytes only become capable of degrading ARE containing mRNAs after the mid blastula stage (175). Figure 3.12 addresses the possibility that this could result from induction or activation of PMR-1 by looking at both changes in expression and phosphorylation at different developmental stages. Figure 3.12A shows the

1 0 2 Figure 3.11 CIAP treatment of PMR-1 alters its mobility on SDS-PAGE. A. 10 pg of partially purified PMR - 1 isolated from the liver 0, 9, or 24 hours after estrogen injection was incubated with 25 U of CIAP for 30 min at 37°C and electrophoresed on a 1 0 % SDS-PAGE gel. And visualized by Westem blot using a polyclonal PMR-1 antibody (ABg). B. 5 pg of baculovirus expressed protein was incubated with 25 U of CIAP for 30 min at 37°. Electrophoresed on a 10% SDS-PAGE gel, and visualized by Westem blot using a polyclonal PMR-1 antibody (ABg). 0. 10 pg of PMR-1 was treated with either 25 U of CIAP (lane 3) or 25 U of CIAP that had been heat deactivated (lane 2) and incubated 30 min at 37°C, and electrophoresed on a 10% SDS-PAGE gel. This was visualized by Westem blot using a polyclonal PMR - 1 antibody (ABg).

103 0 hr 9 hr 24 hr CIAP

Figure 3.11 CIAP treatment of PMR-1 and Baculovirus expressed PMR-60 leads to an altered mobility on SDS-PAGE.

104 Figure 3.12 Development has no effect on the ability of CIAP treatment to alter the mobility of PMR-1 on SDS-PAGE. A. 10 or 20 pg of protein isolated from oocytes and oocytes matured by progesterone was treated with 25 U of CIAP for 30 min at 37°C and electrophoresed on a 10% SDS-PAGE gel. Products were visualized by Westem blot with a polyclonal PMR-1 antibody AB 2 B. 10 pg of protein isolated from stage 3, 9, or 17 embryos was incubated with 25 U of CIAP for 30 min at 37°C. Electrophoresed on a 10% SDS-PAGE gel, transferred to an Imobilon P membrane, and visualized by Westem blot using a polyclonal PMR-1 antibody (AB2 ).

105 / / « y 10 ug 20 ug CIAP - + - + - + - +

1 2 3 4 5 6 7 8

B Stage 3 Stage 9 Stage 17Stage CIAP

Figure 3.12 Development has no effect on the ability of CIAP treatment to alter the mobility of PMR-I on SDS-PAGE..

106 effect of CIAP on protein from stage 3 (4-cell), stage 9 (mid blastula transition), and stage 17 (neurula) embryos. There was no noticeable difference in either the amount of PMR-1 expressed or the effect of CIAP treatment at any of these stages. The experiment in Figure 3.12B shows the effects of oocyte maturation on the expression and phosphorylation of PMR-1. Although there may be a slight induction in the amount of PMR-1 expressed in mature oocytes, there was still no change in the effect of CIAP. This suggests that phosphorylation of PMR-

1 is consistent throughout development, as well as with estrogen stimulation. In vitro activity assays have demonstrated that CIAP treated PMR-1 still maintains its normal activity (Dompenciel and Schoenberg personal communication).

These data suggest that serine/threonine phosphorylation of PMR-1 does not alter its ability to cleave mRNA in vitro. Expression levels and post translational modification of PMR-1 seem to be constant through development and estrogen activation; corroborating the idea that PMR-1 activity is regulated by its interaction with other proteins (Cunningham and Schoenberg submitted).

3.4 Development of LM-PGR for mapping of in vivo degradation intermediates.

3.4.1 A sensitive LM-PGR assay for detectingin vivo mRNA degradation intermediates.

Earlier work found that degradation intermediates generated by in vitro

PMR-1 cleavage of albumin mRNA contained free 3’-hydroxyls, making them susceptible to degradation by 3’-5’ exonucleases (169). This presented an

107 opportunity to develop a method to tag and amplify these fragments by ligating a

primer modified with a 3’-amine group (to prevent concatamerization) coupled to

RT-PCR amplification of the tagged RNA. The protocol described below does

not distinguish between endonuclease cleavage intermediates and pausing of a

3’-5’ exonuclease. However, the latter tend to be highly processive enzymes in vertebrates that are not even retarded by poly(G) tracts (A.-B. Shyu, personal

communication). The LM-PCR protocol is outlined in Figure 3.13. Total RNA is

used, since poly(A) selection would remove the upstream 5’ degradation

intermediates that are detected by this assay. The ligation reaction utilizes a

DNA primer bearing a 5’ phosphate and 3' amino terminus (MHIINH 3 P in

materials and methods), and reverse transcription is primed with a primer, MH12, which is complementary to the modified ligation primer. The cDNA is PCR amplified using a 5’ [^^P]labeled gene-specific primer and the complementary primer (MH12) used for reverse transcription. The products are separated on a denaturing polyacrylamide/urea gel, and bands identified by autoradiography are excised, reamplified, and sequenced. The junction between the ligated primer and the target mRNA sequence identifies the 3’ end of the degradation intermediate.

Efficient ligation of the 5’-phosphate, 3’amino terminal modified primer

MH11NH3 P is essential for success in the later amplification steps. The experiment in Figure 3.14 compares the ligation efficiency for reactions using

108 Fig. 3.13 The LM-PCR protocol for identification of mRNA decay intermediates. The LM-PCR protocol Is based on the observation that PMR-1 decay intermediates have 3’ hydroxyls. A DNA ligation primer bearing a 5’ phosphate and a 3’ amino group is ligated to a preparation of total RNA using RNA ligase. Reverse transcription is primed with a complementary primer, and the resulting cDNA is amplified using the complementary primer and a 5’-end labeled primer specific to the mRNA under study. Amplified products are separated in a denaturing polyacrylamide gel, identified by autoradiography re-amplified by PGR and sequenced to identify the 3’ end of the degradation intermediate at the junction with the ligated primer.

109 degraded OH RNA OH

OH

ligation NH2 primer RNA ligase

NH, reverse transcribe nested primer

•Separate products on urea/acrylamide gel •Autoradiograph •Excise bands and reamplify •Remove primers, sequence PCR products •3’ end is the junction with the ligated primer

Figure 3.13 the LM-PCR protocol for identification of mRNA decay intermediates.

11 0 either polyethylene glycol and a Tris buffer (PEG, lanes 3-5), or dimethylsulfoxide and a Hepes buffer (DMSO, lanes 6-7). These reactions also contained 10 pg of tRNA as a competitor to mimic conditions expected to be encountered in mixtures containing ribosomal RNA and mRNA degradation fragments. Ligation efficiency was determined by a shift in the mobility of a uniformly [^^P]labeled transcript corresponding to the 5’ end of the albumin mRNA (from pXa 470,

(129)), when ligated to the unlabeled primer. The results were quantified by phosphorimager. The percent shifted is indicated below each lane. Polyethylene glycol was significantly more effective than DMSO in promoting primer ligation.

In reactions performed at 18 °C, 37% of the input RNA became ligated to the primer in buffer containing PEG (lane 2) versus only 3.7% in buffer containing

DMSO (lane 7). If both DMSO and PEG were added to either buffer, the results were indistinguishable from PEG alone (compare lanes 4 and 6 with lane 3), and slightly better ligation efficiency (45%) was observed if the reaction was performed at 4°C (lane 5). Based on these results, all subsequent ligation reactions were performed at 4°C using Tris buffer containing polyethylene glycol.

3.4.2 Identification ofin vivo albumin mRNA decay intermediates corresponding to sites cleavedin vitro by PMR-1.

Male Xenopus were injected with 1 mg of estradiol and total liver RNA isolated 0, 12, or 24 hours later was analyzed by LM-PGR for cleavage within the element in the albumin mRNA 5’ coding region. The albumin mRNA 5’ coding regions in vitro cleavage by PMR-1 has been extensively characterized

111 Figure 3.14 Optimization of the ligation protocol. Uniformly [^^P]-labeled 470 nt albumin transcript and 10 [jg of tRNA (lane 1) were ligated to MH 1 1 NH3 P using the PEG ligation method as described in the materials and methods (lane 2). This was also done using several different modifications including addition of DMSO (lane 3), and incubation at 4°C (lane 4). The DMSO ligation method was also used with (lane 6 ) and without addition of 25% (w/v) PEG (lane 5). Samples were separated on a 6 % polyacrylamide/urea gel and visualized by autoradiography. The percent of the transcript that was ligated to MH11NH3P is shown at the bottom as determined using imagequant

112 (0o lUO s 9 p 3 8 o O o (9 w w S U1 UJ s z a. a. o. QQ

Ligated transcript

Transcript

Lane #123 4567 % shifted N/A 0 37 39 45 40 3.7

Figure 3.14 Optimization of the ligation protocoi.

113 (169,139). RNA isolated from lung, which does not express albumin, was used as a negative control. Figure 3.ISA shows the gel separation of RT-PCR products generated by amplification with 5’-[^^P]labeled SetAI primer (which begins at position 39 of albumin mRNA). The numbers on the right side indicate the cleavage positions mapped to the structure of the corresponding region of albumin mRNA in Figure 3.150. A non-specific product that did not come from albumin mRNA was seen to varying degrees in all samples (closed circle). Two major amplification products from time 0 liver RNA were seen in the lower third of the gel; one of which corresponded to cleavage at position 14, and the other of which was too close to the sequencing primer to be identified (open circle). Of note was the time-dependent appearance of new products 12 and 24 hr after estrogen administration (lanes 3 and 4). A doublet product observed in the center of the gel in liver RNA from 24 hr estrogen treated frogs was particularly striking as it resembled the predominant doublet cleavage product generated in vitro by PMR-1 cleavage. These cleavage sites are within the overlapping

APyUGA elements in the mapped stem-loop structure of this portion of albumin mRNA (Fig 3.1 SC) (169).

Each of the indicated bands was excised from the gel, amplified, and sequenced to identify the junction between the ligated primer and albumin mRNA degradation products. An example of this is shown in Figure 3.1 SB for band A11, the strongest of the aforementioned doublet amplification products. The sequence, S’-AUU-primer-3’, corresponded to cleavage within the first APyUGA

114 Fig. 3.15 LM-PCR Identification of decay intermediates in the 5’ coding region of albumin mRNA. A. LM-PCR was performed with 10 p.g of total lung RNA (lane 1) or 10 |ig of total liver RNA prepared 0, 12, or 24 hr after estrogen administration using a 5’-[32P]labeled primer complimentary to a portion of the 5’ end of the mRNA. The PCR products were separated on a 6% acrylamide/urea gel and visualized by autoradiography. The numbers on the right side of the autoradiogram correspond to the sites shown in C. B. The most prominent band (11) was excised, reamplified and sequenced to identify the junction between the ligated primer and the mRNA degradation intermediate. 0. Each of the numbered bands in A was re-amplified and sequenced as in B, and their position is shown on the mapped secondary structure of the 5’ coding region of albumin mRNA bearing the overlapping APyrUGA elements whose in vitro cleavage by PMR-1 was previously characterized (131). The open arrows correspond to in vivo degradation intermediates that were not observed by in vitro cleavage with purified PMR-1.

115 sü

A l A2 m A primer

A6 A7 A albumin C m RNA

A13 A14 A15

o

band A11 sequence

1 2 3 4

A ll

AiO \ r i A ^ccu ^ c C GGA . A A A A

A®A C C AU §H<^A8

(Sa u a c a g o Scaagcauauüg S u a u u g f a- A ïs A13 A12

Figure 3.15 LM-PCR identification of decay intermediates in the 5’ coding region of aibumin mRNA.

116 element in the stem-loop. The positions of the in vivo cleavage sites identified in

this portion of albumin mRNA are indicated in Figure 3.15C with the numbered

arrows corresponding to the LM-PCR products in Figure 3.15A. While many of

these sites correspond to cleavages generated in vitro using an albumin mRNA

substrate transcript and purified PMR-1 (filled arrows), an equal number do not.

There are several possibilities that could account for this difference in

activity between crude extracts and purified PMR-1. One possibility is another

endonuclease present in the crude extracts also cleaves the transcript.

Alternatively, proteins in the crude extracts could interact with either PMR-1 or

the albumin transcript in a way that enhances the ability of PMR-1 to cleave at

non-consensus sites. The latter possibility is supported by an experiment where

100 or 200 U of purified PMR-1 is incubated with total RNA for 30 minutes before

primer extension is performed. This leads to an increased intensity of bands

unique to PMR-1 cleavage, thereby differentiating them from bands due to either

cleavage of another endonuclease or polymerase pausing (Fig. 1.1C, compare

lanes 7 and 8). The eight bands identified by LM-PCR all increased in intensity

when purified PMR-1 was added back to crude extracts from Xenopus liver. This

data suggests that all of the sites mapped are due to PMR-1 cleavage.

These results are the first demonstration of in vivo decay intermediates

corresponding to those generated in vitro by an identified mRNA endonuclease.

The time dependent appearance of these degradation intermediates supports the

117 concept that the 22-fold estrogen induced Increase in unit activity of polysome- bound PMR-1 (Cunningham and Schoenberg, submitted) is the key step in the activation of serum protein mRNA decay. Finally, the cleavage site numbering used here differs from that used in our previous in vitro work (169). This was done to provide a nomenclature consistent with the number of identified in vivo degradation intermediates. In addition, the absolute identification of cleavage sites afforded by the sequencing of LM-PCR products indicated that those reported previously from our in v/fro studies (169) were misplaced by a single nucleotide. This likely results from the use of DNA markers to size RNA degradation intermediates in our earlier work. The correct cleavage sites are indicated in all of the figures presented here.

3.4.3 Application of LM-PCR to identify degradation intermediates from the

3’ end of albumin mRNA.

There are 14 APyUGA elements dispersed across albumin mRNA, many of which are in the 3’ half of the molecule. However, metastable degradation intermediates from this portion of albumin mRNA were not seen in in vitro degradation experiments using uniformly-labeled transcript and polysome extracts from estrogen treated frogs (154). The experiment in Figure 3.16 identified sites of in vivo cleavage in the region spanning from 1690 to the 3’ end of the mRNA using RNA isolated from a 24 hour estrogen treated frog. Eight in vivo cleavage sites were identified in the gel in Figure 3.16A, and their positions in the sequence of the albumin mRNA 3’ end are indicated in Figure 3.16B.

118 Fig. 3.16 LM-PCR identification of decay intermediates in the 3' end of albumin mRNA. A. LM-PCR was performed with 2 pg of total liver RNA from a 24 hr estrogen-treated frog using a primer that amplifies from position 1690 to the end of albumin mRNA. The PCR products were separated on a 6% acrylamide/urea gel and the bands of interest, shown by the arrows, were extracted from the gel and sequenced. B. The sequence of the 3' end of albumin mRNA corresponding to the region analyzed in A is shown with the in vivo degradation intermediates indicated with arrows. Consensus APyrUGA sites are identified with open arrows.

119 B ■Gi

TCCTTGTGAAGCTGATTAAAGTTAGTCCTAAATTGGAA G 8 G7 G6 I $ AAAAATCACATTGATGAATGTTCTGCTGAATTCCTTAA G5 G4 4 150 GATGGTACAGAAATGCTGTACTGCAGATGAACACCAG .0 4 •0 5 G 3 G 2

•06 CCATGTTTTGATACAGAGAAACCAGTACTGATTGAAC G1 100

< = 0 7 ACTGTCAAAAACTCC....JVAA„ ^ 0 8

Figure 3.16 LM-PCR identification of decay intermediates in the 3’end of albumin mRNA.

120 There are 4 APyUGA elements in this sequence, 3 of which could be resolved on the gel. Cleavage site G1 corresponds to the 3’ end of albumin mRNA, and sites

G2, G3, and G7 represent in vivo cleavage within the 3 APyUGA elements

(underlined in Fig 3.16B), None of the cleavages within the APyUGA elements were as strong as the cleavage at non-consensus site G8. These results raised the question of whether there was something intrinsically different about this region of albumin mRNA that effected its ability to be cleaved by PMR-1.

To address this, the in vitro degradation of the 3’ end of albumin mRNA was evaluated using a 5’ end labeled transcript in a manner similar to that used previously to characterize cleavage of the element in the 5’ coding region (169).

In the experiment shown in Figure 3.17A, the 5’ end labeled transcript was incubated with a crude polysome extract from estrogen treated frogs and bands corresponding to all of the in vivo cleavage sites from Figure 3.ISA are labeled on the right. For the most part the pattern observed upon in vitro degradation with the crude polysome extract recapitulated that observed in vivo by LM-PCR.

This sequence contains two overlapping APyUGA elements much like the predominant 5’ coding region PMR-1 cleavage sites, but unlike the 5’ coding region, cleavage at these overlapping elements was minimal (sites G2 and G3).

PMR-1 is a minor component of the extract used in the experiment in Figure

3.17A, and to evaluate its contribution to this pattem the experiment was repeated in Figure 3.17B using 20 units of the purified enzyme. Unexpectedly, only two major cleavages were observed. The first of these is observed at site 7,

121 Fig. 3.17 In vitro cleavage of the 3’ end of albumin mRNA. A.A 5’-end labeled transcript corresponding the 3’ 310 nt of albumin mRNA analyzed In Fig. 4 was Incubated at 23°C for the Indicated times with 10 pg of polysome extract from 24 hr estrogen-treated frogs prepared as described previously (117). The products were separated on a 6% polyacrylamlde/urea gel and visualized by autoradiography, and the degradation fragments corresponding to the in vivo cleavage products are Indicated with arrows. As In Fig. 4, the APyrUGA elements are Identified by open arrows. B. The transcript used In A was Incubated for the Indicated times with 20 U of purified PMR-1. One unit of PMR- 1 Is the amount needed to completely cleave 7 fmol of albumin substrate transcript In 30 mln at 23°C. The products were separated as described In A and are labeled corresponding to the in vivo cleavage sites on the right.

122 B ÿ Time (mln) Time (min) to 2.5 7.5 15 _ 5 20 45 S 0.5 5 10 0 10 30

2 0 0 - 200

1 5 0 - •4 1 5 0 - •5

•6 100

< î = 7 100 — 8

50— — , 1 2 3 4 5 6 7 8 1 2 3 4 5 6

Figure 3.17 In vitro cleavage of the 3' end of albumin mRNA.

123 which is within a consensus APyUGA element, and the other at the adjacent site

8, which is not. in contrast to the S' coding region cleavage sites, there was no detectable cleavage within the overlapping APyUGA element at sites G2 and G3.

This leads us to conclude that some feature of this RNA was interfering with the ability of PMR-1 to cleave within the overlapping APyUGA elements. The lack of cleavage at sites G4, G5, and G6 could either be due to the purification away from another nuclease, or the inability of PMR-1 to cleave these sites due to the loss of an interacting factor.

3.4.4 Impact of secondary structure on PMR-1 cleavage of the albumin mRNA 3’ end.

Because PMR-1 can only cleave single stranded RNA (151,129,169) the most likely explanation for the results in Figure 3.17 is that the overlapping

APyUGA elements in this portion of albumin mRNA are in a structure that is unfavorable for endonuclease cleavage. To test this, the S' end labeled transcript was subjected to limited digestion with RNase T1, and the identified single stranded G residues were used to fit this sequence to a secondary structure using the M-fold program (167). The results of the T1 digestion are shown in Figures.ISA, and the folded structure derived from these results is shown in Figure 3.188. In this structure, the RNase T1 cleavage sites are identified as 'H', the APyUGA elements are boxed, and the in vivo cleavage sites are indicated with arrows. The results indicate that both the APyUGA element at site G7 and the adjacent non-consensus PMR-1 cleavage site G8 are in a large

124 Fig. 3.18 Secondary structure of the3’ end of albumin mRNA. A. The 5’ [^^P]labeled transcript for the albumin mRNA 3’ end was incubated 5 units of RNase T1 and the products separated on a denaturing 6% polyacrylamide gel. The positions of cleaved G residues (identified by dots on the autoradiogram) were determined by mobility relative to size standards. B. The positions of the RNase T1 cleavage sites were used to guide the generation of a predicted secondary structure using MFOLD. The RNase T1 cleavage sites appear as H in the structure, and the 8 cleavage sites mapped in vivo and in vitro are labeled by the arrows. The four APyUGA sequences present are identified by the boxes. Note that the fourth APyrUGA site present in the bottom left portion of the structure was not resolved in the gels used in Figs. 5-7.

125 A i l RNase T1

m / 250 - 1 / h150 // .• 200 - B / 8 / . ^ 'r ' ÿ_ i J % ### 150 - Oo 'VuCc y?" u H \S-- \ AU • i • *uu*u 4*^“ °

# - 1 0 0 100 - ■ - -~ f t

# -A';: y . u ^«CUO- C u o c A , H*

# #

#

50 - - # a fe # ; -5 0 1 2

Figure 3.18 Secondary structure of the 3’ end of albumin mRNA.

126 single stranded bulge. On the other hand, the overlapping APyUGA sites at positions G2 and G3 are in a constrained hairpin structure that would prevent cleavage by PMR-1. This is similar to previous work showing that PMR-1 was unable to cleave within APyUGA when the 5’ coding region element was mutated to form a hairpin structure (169). This could account for the minimal cleavage observed in vivo in figure 3.16 and in vitro using polysome extract in Figure

3.17A. As noted above, a fourth consensus site was present in this transcript, but its location was beyond the resolving power of the gels used in Figure 3.17.

However, this element too was structurally constrained and unlikely to be cleaved in vitro by PMR-1. Taken together, the results in Figures 3.16-3.18 indicate structural constraints limit the ability of PMR-1 to cleave the 3’ end of albumin mRNA both in vivo and in vitro.

3.4.5 Identification of PMR-1 cleavages within the vigilin binding domain of vitellogenin mRNA.

Vitellogenin mRNA, which is induced and stabilized by estrogen in

Xenopus liver with a half-life of over 500 hr, is destabilized by estrogen withdrawal. However even under estrogen withdrawal, it still has a half-life of 15 hours. PMR-1 has been shown in vitro to cleave vitellogenin mRNA at two

APyUGA sites that are part of the binding site of the estrogen induced protein, vigilin (139). The experiment in Figure 3.19 demonstrates that these APyUGA elements are also cleaved in vivo. Ten major bands were identified and sequenced. The two most prominent bands were found (by a gene bank search) 127 Figure 3.19 LM-PCR of vitellogenin B1 mRNA from Xenopus laevis liver. A. LM-PCR was performed with 2 pg of total liver RNA Isolated from primary hepatocytes that had been treated with 10 ® M moxestrol for 5 days, switched to 10'^ M moxestrol for 12 hours prior to t=0 to initiate withdrawal, or 60 hours after withdrawal (t=60). The PCR products were then separated on a 6% acrylamide/urea gel. The closed arrows show the vitellogenin bands that were sequenced and the open arrows show non vitellogenin bands that were sequenced. B. sequence of the vitellogenin B1 3’ UTR. All of the mapped cleavage sites are shown. The APyUGA sites that have been shown to be within the vigilin binding site are underlined.

1 2 8 t î L B

AGCACTGAAATGTGCAGCATGATCCACCCATACCCACATT 8 7 : 1 ACAGGAAGCAGTTGACAATATCTCATATCTCATCAAATGAA BKGRO 6 4 m i i i TAAGCTGAATTCACTGATGATGATAAACTGATCTCAATTTC

BKGRO ACAAACCAAATGTATATTATACTATTGTAAACAATTCAATT 3 2 1 t I I TAAATAAATATTTGCATCCACA(A„)

12 34 5

Figure 3.19 LM-PCR identification of decay intermediates in the viteiiogenin 3' UTR following estrogen withdrawal

129 to be from unknown gene products and are probably due to non-specific binding of the primer to another gene. The other eight bands were sequenced and found to represent vitellogenin degradation intermediates. Bands 5 and 6 were found to represent cleavage at the two APyUGA sites. The relative weak signal of these bands is believed to be due to the continued binding of vigilin to the 3’ UTR of the vitellogenin transcript, which could also account for the relatively long half- life (15 hr) of the ‘unstable’ vitellogenin. This continued association of vigilin with the mRNA could also account for the accumulation of groups of bands like those seen at sites 2 and 8, since vigilin binding could be blocking the normally very processive 3’-5’ exonucleases.

3.4.6 Identification of decay intermediates consistent within vivo endonuclease cleavage within thec-myc CRD.

Ross and co-workers identified an mRNA instability determinant within the coding region of c-myc mRNA (the coding region determinant, CRD, (176)) that is both a site for in vitro cleavage by a polysome associated endonuclease activity (177) and for binding by a KH-domain protein (50). Because little was known about the relationship between the CRD and the degradation of c-myc in vivo, we chose to examine cleavage within the CRD as a test of the ability of LM-

PCR to detect labile in vivo decay intermediates from an inherently unstable mRNA. In the experiment shown in Figure 3.20, LM-PCR was performed on 2 pg of total RNA isolated from murine erythroleukemia (MEL) cells using a gene specific primer complementary to nucleotides 1669-1687 of c-myc mRNA. A 130 Fig. 3.20 LM-PCR mapping of in vitro degradation intermediates within the c-myc coding region determinant. A.LM-PCR was performed as in Fig. 3.15 with 2 pg of total RNA isolated for MEL cells using a primer specific to a region upstream of the c-myc CRD. The PGR products were separated on a 6% acrylamide/urea gel and the bands of interest, shown by the arrows, were excised out and sequenced. B. The sequence of the c-myc CRD is shown with the arrows indicating the positions of the in vivo degradation intermediates identified in A. The underlined region corresponds to the region previously shown to be cleaved in vitro by a polysome-associated endonuclease (135).

131 M c-myc

{ GACCAGATCCCTGAATTGGAAAACAACGAAAAGGCCi 3 2 300 CCCAAGGTAGTGATCCTCAAAAAAGCCACCGCCTAC 250

200 - f ATCCTGTCCATTCAAGCAGACGAGCACAAGCTCACCT ■2 1 150 — • 3 II CTGAAA^GACTTATTGAGGAAACGACGAGAACAGTT

.5 GAAACACAAACTCGAACAGCTTCGAAACTCTGGTGCA

100

1 2

3.20 LM-PCR identification of degradation intermediates within the c-myc coding region determinant.

132 denaturing polyacrylamide gel of the [^^PJIabeled LM-PCR products Is shown In

Figure 3.20A, and the sequence of this portion of c-myc mRNA showing the locations of the 5 Identified in vivo mRNA degradation Intermediates Is found In

Figure 3.20B. As with albumin mRNA, each of these sites was determined by the sequence of the junction between the ligated primer and the c-myc mRNA degradation Intermediate. Band 4 Is particularly noteworthy here as this corresponds to a previously mapped site for in vitro cleavage of c-myc mRNA

(177). Thus, this supports the notion that the In vivo degradation of c-myc mRNA

Involves endonucleolytic cleavage. In addition, these results demonstrate that the LM-PCR approach to mapping in vivo mRNA decay Intermediates Is generally applicable to both highly abundant mRNAs, like albumin, and rare mRNAs, like c-myc.

3.5 PMR-1 antisense Morpholino has no effect on level of albumin mRNA in primary hepatocytes.

Morpholino oligonucleotides are a new type of antisense ollgo designed by

Gene Tools, LLC. (Corvallis,OR). They have a modified backbone, with an amino group replacing one of the oxygens In the phosphate group, which prevents rapid degradation In the cell. This Is Important since many antisense ollgos are degraded before they have a chance to significantly effect the expression of their target gene. One drawback of this modification Is that It creates an uncharged backbone which makes It a poor substrate for cell culture transfection using standard llpid reagents. This was addressed by annealing the

133 Morpholino ollgo, Std-control, or AS-PMR to a complementary DNA ollgo containing a short poly(A) tall, MH32 or MH33, respectively. This duplex can then be transfected Into cells using standard llpid reagents (see materials and methods). In theory, the DNA ollgo will be rapidly degraded by the cell, while the

Morpholino will remain stable.

Table 3.1 shows the transfection efficiency observed when using between

0.025 pM and 1.0 pM of Morpholino transfected Into primary hepatocytes using either Effectene™ or LIpofectamlne™ transfection reagenis. The

LIpofectamlne™ reagent, at Its best, was only able to transfect about 8% of the primary hepatocytes. It also caused a significant decrease In the viability of the cells. On the other hand, between 0.025 pM and 0.3 pM o f the Morpholino transfected with Effectene™ reagent was able to transfect approximately 85% of the hepatocytes and did not cause a significant decrease an cell viability. Figure

3.21 shows an example of the transfection efficiency of 0.05 pM of the fluorescelnated Std-control Morpholino 2 hr after being transfected Into primary hepatocytes using the Effectene™ reagent from QIagen (Valencia, CA) and visualized by confocal microscopy using either a phase lamp (top) or fluorescent lamp (bottom).

Based on the high transfection efficiency above 0.0-5 pM of AS-PMR, an antisense PMR-1 Morpholino, or the Std-control ollgo, was transfected Into primary hepatocytes using Effectene™ reagent. Twenty four hours later cells 134 Figure 3.21 Transfection efficiency of Morpholino oligo into primary hepatocytes. Primary hepatocytes were transfected with 0.05 pM of a control 3’-fluoroscein labeled standard control Morpholino oligo. Two hours latter the cells were washed with PBS + glucose and visualized by confocal microscopy using either halogen lamp with no filters (top), or a mercury bumer with the IB dichroic mirror to detect the fluorescein labeled Morpholino (bottom).

135 Phase

Florescent

Figure 3.21 Transfection efficiency of Morpholino oiigonucleotieotides with primary hepatocytes

136 Transfection reagent/ Percent of cells concentration of morpholino transfected

Lipofectamine (all concentrations) <8% Effectene 0.0025 uM 85% Effectene 0.05 uM 87% Effectene 0.1 uM 89% Effectene 0.2 uM 87% Effectene 0.3 uM 87% Effectene 0.4 uM 55% Effectene 0.5 uM 0% Effectene 0.8 uM 0% Effectene 1.0 uM 0%

Table 3.1 Transfection efficiency of Morpholino oligo into primary hepatocytes.

137 Figure 3.22 Northern blot of total hepatocyte RNA from Morpholino treated cells. Total liver RNA Isolated from hepatocytes that had been transfected with 0.05 pM of AS-PMR or a standard control Morpholino 48 hours prior was electrophoresed on a 1% agarose gel and transferred to a nitrocellulose membrane. This was probed with 10,000 cpm of random primed albumin or ferittin probes and the amount of albumin mRNA present was normalized to the ferritin mRNA and shown under each lane.

138 9

Estrogen

■ Albumin

Ferritin

1 2 3 4 5 6 % Albumin 97 55 98 55 100 46

3.22 Northern blot of total hepatocyte RNA from cells transfected with Morpholino antisense ollgos.

139 were treated with 1 X 10'^ M moxestroi for 48 hours in order to induce PMR-1 activity. Figure 3.22 shows a northern blot of cytoplasmic RNA isolated from these cells. The percent of albumin mRNA normalized to feritin is shown below each lane. Neither the Std-control nor the AS-PMR Morpholinos had any effect on the expression of albumin mRNA, with approximately 50% of the transcript being degraded upon estrogen induction in all cases. There are several factors that could explain this. One possibility is that PMR-1 does not affect the stability of albumin mRNA in an estrogen dependent manner. However, PMR-1 has been well documented to be involved in the estrogen induced degradation of albumin

(129,169,156,139). More likely, the half-life of PMR-1 is long enough that it is not cleared from the cells in 48 hours, or PMR-1 expression is not blocked by the antisense oligo used. Unfortunately, enough protein could not be isolated from the cells to be able to properly address this by looking directly at the levels of

PMR-1 expression.

140 CHAPTER 4

DISCUSSION

4.1 PMR-1 and hMPO are biochemically different from each other.

PMR-1 is a ribonuclease whose activity appears on Xenopus liver

polysomes concomitant with the estrogen-induced destabilization of mRNAs for

the major serum proteins (154). The identity of cleavage products generated in

vivo following estrogen treatment with those generated in vitro by the purified

enzyme indicates that PMR-1 is a messenger RNase whose activation/induction

by estrogen is an (or the) initiating event in mRNA destabilization. PMR-1 was

purified (129), and its cDNA was cloned (156). Perhaps the most striking

observation to come of this effort was the discovery that PMR-1 is unrelated to

any of the known vertebrate ribonucleases, but instead is a member of the

peroxidase gene family. Given the very high sequence identity (57%) between

PMR-1 and hMPO and the evolutionary distance between Xenopus and humans

one might assume that the protein whose cDNA we cloned was frog

myeloperoxidase. In fact, PMR-1 shares several characteristics with hMPO,

such as its synthesis as a larger precursor that is proteolytically processed to the final product. However, several observations suggested otherwise. First,

myeloperoxidase is a 150 kDa heterotetramer consisting of two light and two

141 heavy chains, both of which are processed from the larger precursor, whereas

PMR-1 is only ever seen as a -60 kDa monomeric protein. Second, hMPO is

found in polymorphonuclear leukocytes, whereas PMR-1 was purified from liver

and its cDNA cloned from the same source. Third, hMPO has a hydrophobic

signal peptide which is lacking from PMR-1 (Fig. 3.1). These differences led us

to examine here in more detail the biochemical relationship between PMR-1 and

the peroxidases, particularly its closest relative hMPO.

The post-translational processing of hMPO in the endoplasmic reticulum

and Golgi involves proteolysis processing and the addition of both heme and

complex mannose N-linked oligosaccharides (178,179,158). Heme addition is a

particularly important step, as it both precedes, and is required for proteolysis of

the precursor peptide into light and heavy chains (178). The data in Figure 3.2

and 3.3 indicate that PMR-1 does not contain covalently-bound heme. Since

heme addition occurs during processing in the endoplasmic reticulum, this finding

is consistent with the absence of a hydrophobic signal peptide on PMR-1. The

data in Figures 3.5 and 3.6 indicate PMR-1 lacks the N-linked glycosyiation

characteristic of hMPO. Once again, these results are consistent with the

absence of post-translational processing of PMR-1 within the endoplasmic

reticulum and Golgi. Nevertheless, PMR-1 does undergo post-translational

processing. The sequence of PMR-1 cDNA indicates it is made from a larger precursor, and the predicted 80 kDa pro-PMR-1 has been detected on prolonged exposure of Westem blots of hepatocyte protein.

142 Interestingly, although PMR-1 and hMPO have distinct biochemical activities they seem to share an . The heme in hMPO is coordinated by two histidines at positions 261 and 502. These amino acids are essential for the peroxidase activity of hMPO (161). Preliminary data suggests that if both of these histidines are mutated to alanine that PMR-1 activity is lost (Peng and

Schoenberg personal communication).

These data suggest that PMR-1 is a novel member of the peroxidase family that has evolved a unique activity as an RNA endonuclease. Despite this unique activity the sequence similarity suggests that the structure of PMR-1 can be predicted based on the crystal structure of canine MPO (180). Using the coordinates from this structure as a guide we were able to create a predicted structure for PMR-1 using Swiss-pro, which is shown in figure 4.1. The most striking feature of this structure is the presence of a grove that is 19 angstroms wide, shown by the two arrows. This is just the right size to fit a RNA molecule, and would make a reasonable active site with RNA cleavage being coordinated by the two histidines, shown in green. Interestingly in MPO these histidines are involved in coordinating the heme group, and are therefore integral to its activity.

The coordination of RNA cleavage by histidines can be seen in RNase A, which has been demonstrated to use histidines to coordinate the cleavage of RNA

(181).

143 Figure 4.1 Predicted structure of PMR-1. The coordinates of canine MPO were used with swiss-pro to determine the most likely structure of PMR-1. The arrows shows a 19 Â deaf that could be a RNA binding region and the two histidines predicted to coordinate the cleavage reaction are shown in green.

144 f

%-%' ; 5 -’ ■ ■ ■ '' # # / ' ' ' 1^® % i ,ÿ-

, - £ , % a # ^

' Y:A \. h \

1V ->p ■

Figure 4.1 Predicted structure of PMR-1.

145 Despite the presence of this potential RNA binding pocket, the sequence of PMR-1 does not have a known RNA binding domain. However, PMR-1 has been demonstrated to be part of a large multiprotein complex suggesting that

PMR-1 may interact with other proteins, and these could be involved in targeting

PMR-1 to various RNA substrates (183).

4.2 PMR-1 is expressed in multipleXenopus and murine tissues.

It is widely believed that there are a small number of endonucleases which are involved in the regulated degradation of a large number of transcripts in a tissue and hormone dependent manner in higher eukaryotes (1). However,

PMR-1 is currently the only endonuclease in higher eukaryotes that has been purified to homogeneity and had a cDNA clone identified. Therefore, PMR-1 represents the first opportunity to use antibodies, as well as activity to identify the expression patterns of an endonuclease. By using this dual approach, we were able to show that most if not all Xenopus tissues express a protein of the same size as PMR-1 that also cross reacts with a polyclonal antibody, ABa, which was raised against PMR-1 (Fig.3.8). Corresponding to this, all of these tissues were demonstrated to have an activity similar to PMR-1, producing the characteristic doublet cleavage in standard in vitro activity assays (Fig. 3.9). The oviduct, which was the only tissue that did not seem to have a cross-reacting protein by

Westem blot analysis, was very viscous and by Coomosie stain seemed to have less protein on the membrane. This is believed to be due to most of the viscous sample being retained in the well of the gel. Taken together, these data suggest

146 that PMR-1 is expressed and active in most if not all tissues in Xenopus laevis.

The presence of an active PMR-1 in these tissues suggests a wider function in mRNA stability than just the liver specific degradation of serum protein and vitellogenin mRNAs.

As well as being widely expressed in Xenopus, PMR-1 may also be conserved in mammals based on the presence of an abundant protein in murine tissues that cross reacts with two independent polyclonal antibodies raised against PMR-1 (Fig. 3.8) (data not shown). However, this protein does have a slower mobility than PMR-1 in SDS-PAGE, suggesting that it may be processed differently. One possibility is that it could be proteolytically processed from the

80 kDa precursor protein at a different site leading to the production of a slightly larger active protein. This is supported by the fact that the same decrease in mobility is seen with baculovirus expressed PMR-60. PMR-60, like the murine activity, is unable to replicate the major doublet cleavage, despite being able to replicate the minor cleavages produced by PMR-1 (Fig. 3.10)(156). We were unable to identify the exact N-terminus of PMR-1 by peptide sequencing because it was blocked. This led to a best guess of where that processing may occur in the production of the PMR-60 clone based on the identified processing sites between the heavy and light chains of hMPO.

Interestingly, Kirsten Bremer, has identified a protein in mouse erythroleukemia (MEL) cells that also cross reacts with PMR-1 antibodies and

147 has a decreased mobility on SDS-PAGE when compared to PMR-1. These extracts have a specific activity that degrades (3-globin mRNA and have been shown to produce cleavages in albumin mRNA that are similar to the secondary cleavages produced by PMR-1. Again, the protein is unable to produce the characteristic doublet product (Bremer, K., and Schoenberg, D.R., personal communication). This activity in MEL cells is seen best when partially purified away from the abundance of nuclease activity present in crude extracts. Taken together, these data suggest that PMR-1 is expressed in most murine tissues and may be proteolytically processed at a slightly different site than in Xenopus.

This could lead to the production of a slightly larger protein with a similar but not identical activity to PMR-1.

This wide spread expression of PMR-1 suggest that it might be involved in the degradation of a large number of transcripts in a tissue and environment specific manner. One possibility mentioned above is that altered post translational modifications could alter the specificity of the enzyme. However it is also possible that in place of or in addition to this PMR-1 could form complexes with various proteins that target it to different messages and thereby dictate its specificity. We do know that PMR-1 is present in a large macromolecule complex of over 660 kDa (183). This complex suggests that PMR-1 interacts with other proteins, and any one of these could be involved in targeting PMR-1.

148 4.3 The effect of CIAP treatment does not change with estrogen or development.

PMR-1 has been demonstrated to be activated in an ER-dependent but transcription independent manner in Xenopus liver (153). A growing body of literature demonstrates that estrogen can activate a number of kinases involved in several different signaling pathways (182). This would suggest that PMR-1 might also be activated by estrogen induction of one of these kinases. However, all of the data collected to date suggests that PMR-1 is not tyrosine phosphorylated, and that serine/threonine phosphorylation does not change with estrogen stimulation of PMR-1 activity (Fig. 3.11 and 3.12), and is not essential for PMR-1 activity. Given that PMR-1 is activated by estrogen in a transcription independent manner, but its own phosphorylation is not altered, it is likely that the ER may phosphorylate a protein that interacts with PMR-1, which has been demonstrated to be part of both the mRNP and polysome complexes (183). Also an inhibitory activity has been identified in SI 00 extracts from estrogen treated frogs (154). These data support the idea that PMR-1 may be activated by the release of an inhibitor, whose binding could be regulated by ER phosphorylation.

Since the levels of expression and serine/threonine phosphorylation of PMR-1 itself do not change during development or estrogen activation of its activity in the liver, PMR-1 is most likely activated by a change in phosphorylation of another protein such as an inhibitor. The data to date suggests that upon

149 estrogen stimulation the inhibitor is phosphorylated and this leads to dissociation of the inhibitor from PMR-1, activating the nuclease.

4.4 Development of LM-PCR method.

Whether a particular mRNA is degraded through an endoribonuclease mediated pathway or an exoribonuclease pathway, with very few exceptions mRNA degradation in vivo is not accompanied by the appearance of stable degradation intermediates. The most likely explanation for this is that in vivo these intermediates are subject to rapid clearance by exoribonucleases.

However, decay intermediates have been observed using crude in vitro decay systems. Primer extension and S1 protection assays have been used successfully to identify in vivo endonuclease cleavage of apo-very low density lipoprotein II mRNA (135) and transferrin receptor mRNA (183). We have used both of these approaches to demonstrate in vivo cleavage within two overlapping

APyUGA elements in the 5’ coding region of albumin mRNA (Fig 1.1)(163).

However, there are numerous disadvantages to these techniques. They work best with highly abundant mRNAs, and even then primer extension and SI protection assays can require up to a 30 day exposure to visualize decay intermediates. These long exposure times can make it difficult to differentiate background signals from actual bands.

We have developed a ligation mediated PCR method for the rapid and precise mapping of in vivo mRNA decay intermediates. The advantages of this

150 method include Its ease of implementation and ability to identify degradation

intermediates from even rare mRNAs, such as c-myc. Although LM-PCR is

unable to differentiate between the 3’ end of a decay product generated by

endonuclease cleavage and a pausing site for a 3’-5’ exonuclease, Shyu and co­

workers have tried without success to show in vivo exonuclease pausing using

poly(G) tracts in a manner similar to that used by Parker and co-workers with

yeast (73, personal communication). This suggests that mammalian 3’-5’

exonucleases are highly processive and that secondary structure does little to

slow these enzyme complexes down. From this, the conclusion that the majority

of sites mapped will be due to endonuclease cleavage products that have not yet

been recognized by the exonuclease complex can be drawn.

4.4.1 LM-PCR Identifies the same cleavagesin vivo as previously mapped

in vitro.

Previous work from the lab has identified PMR-1 as an estrogen induced

endonuclease, whose activity appeared on polysomes coincident with the estrogen induced destabilization of albumin and other serum protein mRNAs

(154). PMR-1 was purified based on a characteristic doublet cleavage it produced within two overlapping APyUGA elements in the 5’ portion of albumin mRNA in vitro. This same doublet was identified as a product of the in vivo degradation of albumin mRNA by S I protection assay (163) and primer extension experiments (Fig. 1.1). LM-PCR identified 8 potential cleavage sites within the 5’ portion of albumin mRNA that has been extensively characterized in

151 vitro (169). Four of these sites were Identical to those mapped in vitro. Including

the characteristic doublet, while 3 other sites were also mapped in vivo by primer

extension. The one remaining site corresponded to a weak band and Is believed

to demonstrate the Increased sensitivity of the LM-PCR method compared to

primer extension.

When purified PMR-1 Is added back to extracts that are then used for

primer extension mapping, all 7 of the band mapped in vivo Increased In Intensity

(Fig. 1.1), suggesting that they are all due to PMR-1 cleavage. The presence of

additional cleavage sites in vivo compared to purified enzyme in vitro could be

due to a difference In the secondary structure of the mRNA, and/or PMR-1

Interacting with proteins in vivo that alter Its specificity for cleavage. The

association of PMR-1 with the mRNPs and polysomes supports the Idea that

PMR-1 Interacts with other proteins in vivo.

Unlike the in vitro assays that show cleavage at sites 10 and 11 as being equal In intensity (169), both in v/vo assays, primer extension (Fig. 1.1) and LM-

PCR (Fig. 3.15) demonstrated site 11 as the most Intense cleavage site. Since primer extension maps sites from the 3’ end and LM-PCR maps sites from the 5’ end, this apparent preference for cleavage at site 11 Is not an artifact caused by masking of downstream sites, and Indicates a true preference for cleavage at this site in vivo. This data supports the Idea that either albumin mRNA, PMR-1, or

152 both interact with other factors in vivo which effect the specificity of PMR-1

cleavage.

4.4.2 In vivo mapping of the 3' portion of albumin is consistent with PMR-1

as the major factor involved in the degradation of albumin mRNA.

LM-PCR was also used to map cleavage sites in the 3' end of the albumin

mRNA, where 4 APyUGA sites are located. Surprisingly, only one of these

consensus sites was a major cleavage site, while two others were shown to be

minor cleavage sites. The fourth site was not located within the resolution of the

gel (Fig.3.16). This same preference was seen when crude extract was used to

map cleavage sites in vitro (Fig. 3.17A). Interestingly, when purified PMR-1 was

used to cleave in v/Yro transcripts only the major cleavage was seen (Fig. 3.17B).

However, when RNase T1 mapping was used to identify regions that were single

stranded, and this data was used in M-fold to predict the secondary structure of

the 3’ portion of albumin, the two consensus APyUGA sites that were not cleaved

by the purified enzyme were found to be present in a double stranded structure.

This supports the hypothesis that PMR-1 requires single stranded mRNA for cleavage (Fig.3.18).

The 3’ end of the transcript both in vivo and in crude extracts in vitro mapped the same cleavage sites as each other. However, purified PMR-1 only produced two of the eight cleavages seen in these crude extracts. These data do not rule out the possibility that another nuclease is responsible for the

153 cleavages identified only in the crude extracts. However, when considered in conjunction with the data from the 5’ portion of the transcript (Fig. 1.1 and Fig.

3.15), these data support the idea that PMR-1 or albumin mRNA are interacting with other factors in crude extracts that alter the specificity of PMR-1 cleavage.

The similarity seen between the crude extracts in vitro and the in vivo cleavage sites suggest that crude extracts can, in many instances, be used to accurately determine the cleavages that occur in vivo.

These data also demonstrate the effect of secondary structure on the ability of PMR-1 to cleave mRNA. This suggests that one must always be aware that stabilizing or destabilizing effects seen from deletions or insertions could be due to changes in secondary structure of other parts of a message as apposed to just the presence of an elements within an mRNA. This idea demonstrates the importance of combining loss of function and gain of function experiments before prediction the presence of a functional RNA element.

4.4.3 LM-PCR suggests that PMR-1 is also involved in the degradation of vitellogenin mRNA.

Shapiro’s lab has previously identified a protein, vigilin, that stabilizes vitellogenin mRNA by binding to the 3’ UTR. The binding site was identified and found to contain the only two APyUGA sites identified in vitellogenin mRNA.

Work done in collaboration with Shapiro has demonstrated that vigilin and PMR-1 compete for binding to the vitellogenin 3’ UTR, leading to either stabilization or

154 degradation on the mRNA respectively (139). The data presented here (Fig.

3.19) demonstrates in vivo cleavage of the vitellogenin mRNA at both of these sites in a manner consistent with PMR-1 degradation. Both the relative weak signal of these bands and the production of clusters of bands as apposed to the discrete bands seen with the albumin and c-myc mRNA is believed to be due to the continued binding of vigilin to the 3’ UTR of the vitellogenin transcript interfering with endonuclease cleavage. Given the relatively long half-life (15 hr) of the ‘unstable’ vitellogenin mRNA, this would not be surprising. However, it Is also possible that these groups of bands are the product of a 3'-5’ exonuclease, whose processivity is inhibited by vigilin binding to the mRNA.

4.4.4 LM-PCR can also be applied to rare mRNAs such asc-myc.

Jeff Ross’ lab has previously identified a region of the c-myc mRNA called the coding region determinant that is responsible for its regulated destabilization in a translation dependent manner (184,50,5). They have identified a cleavage site in vitro that occurs in the coding region determinate of c-myc (177). LM-PCR was used to demonstrate that a cleavage within this region also occurs in vivo

(Fig. 3.20). There are also several cleavage sites mapped that do not fall within this short region.

Taken together, these data demonstrate that a new easier method has been developed for the mapping of in vitro cleavage sites that can be applied to even rare mRNAs such as c-myc. This should expedite the task of identifying

155 sites of endonuclease attack that are responsible for th e in vivo degradation of many mRNAs, greatly expanding the understanding o i how mRNA stability is regulated.

4.5 Morpholino antisense PMR-1 oligos have no e~ffect on the estrogen destabilization of albumin in hepatocytes.

An antisense Morpholino oligonucleotide was diesigned by Gene Tool,

LLC. to optimize its ability to block expression of PM R-1. Using a fluorescienated standard control oligo, we were able to demonstrate that approximately 85% of primary hepatocytes are transfected using Effectene^*^ reagent (Fig. 3.21). This system was then used to determine what effect blocking expression of PMR-1 has on the estrogen induced destabilization of albumin using the AS-PMR oligo.

However, no change in the amount of albumin mRNA was evident, even though

PMR-1 has been well characterized as playing an important role in the estrogen induced degradation of albumin mRNA (Fig. 3.22)(139,156,169,129,132).

Several different factors could account for this discrepancy. First, PMR-1 could have a half-life that is long enough to prevent the clearance of the protein from the cell in 48 hours. This would prevent a significzant decrease in the amount of active PMR-1, which would lead to normal levels of degradation.

Unfortunately, the primary hepatocytes are not viable iin culture long enough to increase the time of treatment, nor were we able to iso*late enough protein to identify if the level of PMR-1 expression changed. However, the lab is working

156 on developing a new cell culture system with A6 cells (Xenopus kidney) that

preliminary data suggest mimics the activity seen in Xenopus liver. This system

could be used to address this possibility (Peng.J., Bremer,K.A., and Schoenberg

unpublished). Second, the AS-PMR Morpholino may have been unable to block

the expression of PMR-1. Given that Xenopus laevis has a duplicated genome

and only one copy of PMR-1 has been cloned, it is possible that the two genes

have different sequences at the 5’ terminus. It is possible that this difference in

the duplicated genes could account for the doublet at 62/64 kDa seen on

Western blots. If this is the case, then the AS-PMR Morpholino may not be able to block expression of PMR-1 by itself.

4.6 Concluding remarks

All mRNA have a 5’ cap and 3’ poly(A) tail both of which function to stabilize the mRNA by blocking exonuclease decay. The degradation machinery has to pass through these structures to degrade the mRNA. The cell is able to get past these structures by two different mechanisms. The first was described by Roy Parker in yeast as a general pathway for mRNA turnover. This pathway is initiated by the poly(A) tail being shortened to oligo(A) lengths. The loss of the poly(A) tail triggers removal of the ^"^G cap, and the decapped transcript is then degraded by the 5’-3’ exonuclease Xrnip (73). Although mammalian homologs to most of these genes have now been identified no one has been able to prove that this pathway also occurs in mammals. Despite this it is believed that mammals have a similar pathway for the general turnover of mRNA involving

157 removal of the cap and/or poly(A) tail shortening and exonuclease decay.

Along with this role in general mRNA turnover exonucleases are also involved in the degradation of extraneous RNA such as endonuclease cleavage products.

The second method the cell has to bypass the protective structures is endonuclease decay. Interestingly this seems to be used for the destabilization of specific mRNAs or subsets of cellular mRNAs under specific conditions in certain tissues. Therefore endonuclease decay seems to be targeted while exonuclease decay is general. The data presented in this work as well as other work from the lab suggest that PMR-1 is widely expressed in many organisms and tissues, and is involved in the targeted degradation of a large number of transcripts in a tissue and environment specific manner. I have demonstrated that PMR-1 degrades serum protein mRNAs such as albumin in Xenopus liver in an estrogen dependent manner, and degrades vitellogenin mRNA upon estrogen withdrawal. Preliminary data in the lab shows that an activity identified in MEL cells and found to produce many of the same cleavages as purified PMR-1 in vitro degrades (3-globin mRNA during cell differentiation. I believe that these examples are but the first of many mRNAs that will be found to be degraded by

PMR-1 in a tissue and environment specific manner.

PMR-1 is bound to polysomes in the context of a >660 kDa mRNP complex. In conjunction with the presence of PMR-1 in these complexes the existence of an inhibiting activity in SI GO extracts from estrogen stimulated

158 Xenopus liver suggest that PMR-1 may interact with other proteins. Since PMR-

1 expression and phosphorylation are not altered when PMR-1 is activated in

Xenopus liver, it is most likely that PMR-1 is regulated by its interaction with one or more proteins that can act as an inhibitor when bound to PMR-1. This would be consistent with the regulation of other endonucleases such as the RNase A family and its inhibition by placental ribonuclease inhibitor, and RNase L and its inhibition by RNase L inhibitor (185).

Regulation of endonucleases by inhibitor binding leads to two possible methods for this selective activation. First, different tissues could express different isoforms of the inhibitor either through alternative splicing or expression of different members of a family of proteins. Binding of these isoforms to the endonuclease could be regulated through different signaling pathways such that any particular inhibitor is only affected by one or two signaling pathways.

Second, the same inhibitor could be expressed in all tissues and the presence or absence of certain pathways within certain tissues could regulate the binding of the inhibitor to the endonuclease. Either one of these pathways could account for tissue and environment specific activation of the endonuclease.

The above model is consistent with the fact that mis-expression of an endonuclease would likely be toxic to the cell. The regulation of endonuclease activity by an inhibitor allows for both rapid initiation of decay as well as rapid inhibition of activity since no new protein needs to be produced. This allows for

159 finer control of endonuclease activation and inactivation mediating mRNA decay to prevent the degradation of non-target mRNAs after the target has been largely cleared from the cell. It is important to remember that most endonuclease are sequence selective and not sequence specific. For example although purified

PMR-1 was shown to degrade albumin transcript preferentially over the antisense transcript in vitro, if given enough time, PMR-1 is able to degrade the antisense transcript. Therefore an unregulated endonuclease could eventually degraded ‘non traditional’ targets if left unchecked. This fact could be the most convincing argument for the need of inhibitors to quickly shut down activity after degradation is complete.

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