MECHANISM OF RNA REMODELING BY DEAD-BOX

by

QUANSHENG YANG

Submitted in partial fulfillment of the requirements

For the degree of Doctor of Philosophy

Thesis Advisor: Dr. Eckhard Jankowsky

Department of Biochemistry

CASE WESTERN RESERVE UNIVERSITY

May 2007 CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

______Quansheng Yang______

candidate for the______Ph.D.______degree

(signed)______William Merrick______

(Chair of the Committee)

______Vernon Anderson______

______Eckhard Jankowsky ______

______Anthony Berdis ______

______

______

(date) ______12/12/2006______

2 Table of Contents

Chapter 1: Structures and biochemical activities of DExH/D helicases………………...11 Chapter 2: Initial characterization of Ded1…………….....…………………...... 31 Chapter 3: ATP and ADP-dependent modulation of RNA unwinding and strand annealing activities by the DEAD-box Ded1…………………………………...41 Chapter 4: Protein-assisted RNA structure conversion towards and against thermodynamic equilibrium values………………………………………………………68 Chapter 5: Duplex unwinding by a DEAD-box helicase without translocation on the loading strand...………………………………………………………………...... 101 Chapter 6: Duplex unwinding by DEAD-box helicases from both terminal and internal helical regions…………………………………………………………………………..125 Chapter 7: Future directions……………………………………………….……………146 Chapter 8: Materials and methods………………………………………………...…....151

3 List of Tables

Table 1.1 NTPase activities of DExH/D helicases……………………………………....16 Table 1.2 Polarity of RNA duplex unwinding by DExH/D helicases…………………...18 Table 1.3 Unwinding capability of DExH/D helicases……………………………….....21 Table 1.4 A selection of DEAD-box helicases containing profound annealing activity...26 Table 2.1 Substrates and their sequences………………………………………………...34 Table 3.1 Substrates and their sequences………………………………………………...43 Table 4.1 Substrates and their sequences………………………………………………...72 Table 5.1 Substrates and their sequences used in Figure 5.1…………………………...105 Table 5.2 Sequences and characterization of substrates used in Figure 5.4 …………...110 Table 5.3 Sequences of multi-piece substrates (MPS)………………………………….114

4 List of Figures

Figure 1.1 Structure and sequence characteristic of RNA helicases...…………………..14 Figure 2.1 Ded1 has RNA-dependent ATPase activity and ATP-dependent unwinding activity……………………………………………………………………………………33 Figure 2.2 Effects of the pH and salt concentration on the unwinding activity of Ded1 for 13 bp duplex RNA containing a 25 nt ssRNA at the 3’-end...... ……………...36 Figure 2.3 Effects of the pH and salt concentration on the unwinding activity of Ded1 for 13 bp blunt-end duplex …………………………...... ……………………………...….37 Figure 3.1 RNA unwinding and strand annealing activities of Ded1…………….……...44 Figure 3.2 Pronounced strand annealing activity is specific for Ded1………….……….46 Figure 3.3 Ded1-catalyzed ATP-dependent steady state between RNA unwinding and strand annealing………………………………………………………………………….48 Figure 3.4 Dependence of unwinding and annealing rate constants on ATP-concentration and the nature of the RNA substrate……………………………………………………..50 Figure 3.5 ADP-dependent modulations of unwinding and annealing activities of Ded1...... 57 Figure 3.6 Effects of AMPPNP on unwinding and annealing activities of Ded1………..59

Figure 3.7 MgCl2-dependent modulation of unwinding and annealing activities of Ded1……………………………………………………………………………………...60 Figure 4.1 Substrate design and characterization………………………………………..71 Figure 4.2 Ded1 can unwind complex A and B and anneal complex A and B from respective RNA strands………………………………………………………………….74 Figure 4.3 RNA structure conversions…………………………………………………..75 Figure 4.4 Representative time course of structure conversion A Æ B with 800 nM Ded1 and 0.5 mM AMPPNP…………………………………………………………………...79 Figure 4.5 Basic kinetic scheme for structure conversion with Ded1 and ATP………....81 Figure 4.6 Ded1- assisted stabilization of tripartite intermediate………………………..83 Figure 4.7 Measurement of Ded1- assisted stabilization of tripartite intermediate by single molecule FRET……………………………………………………………………85 Figure 4.8 Basic kinetic schemes for Ded1-assisted structure conversion without ATP..89 Figure 4.9 Possible branch migration intermediates during Ded1-assisted structure conversion without ATP…………………………………………………………………90 Figure 4.10 Coupling of RNA structure conversion to deposition of U1A……………...93 Figure 4.11 Ded1-assisted RNA structure conversion…………………………………...95 Figure 5.1 Ded1 unwinds RNA duplexes irrespective of the orientation of single stranded regions…………………………………………………………………………………..104

5 Figure 5.2 Supplementing single stranded RNA in trans does not enhance the basal unwinding rate constant of the blunt-end substrates…………………………………....106 Figure 5.3 Equilibrium binding of different substrates by Ded1……………………….108 Figure 5.4 Unwinding of RNA-DNA hybrid substrates by Ded1……………………...109 Figure 5.5 Unwinding of multi-piece substrate I (MPS I) by Ded1 but not by NPH- II………………………………………………………………………………………..113 Figure 5.6 Unwinding of multi-piece substrate II by Ded1 but not by NPH-II………..116 Figure 5.7 Unwinding of the multi-piece substrate III by Ded1 but not by the DExH RNA helicase NPH-II…………………………………………………………...... ……….118 Figure 5.8 Unwinding of MPS III components without streptavidin……….………….119 Figure 6.1 Dependence of unwinding rate constants on increasing Ded1 concentrations…………………………………………………………………….…….127 Figure 6.2 Unwinding of RNA-DNA chimeric substrates by Ded1…………………....129 Figure 6.3 Ded1 unwinds duplexes lacking any free RNA terminus…………………..131 Figure 6.4 Unwinding within the helical region by Ded1 and Mss116………………...133 Figure 6.5 Both Ded1 and Mss116 do not preferentially initiate unwinding from the terminus……………………………………………………………………………..…..137 Figure 6.6 The unwinding ability of both Ded1 and Mss116 decreases with the RNA length within the helical region………………………………………………………...140 Figure 6.7 DEAD-box helicases unwind RNA duplexes from both internal and terminal helical regions……………………………………………………...... ……….….142

6 Acknowledgements

I would first like to thank my thesis committees, Drs. William Merrick, Vernon Anderson, and Anthony Berdis. I thank them for their very useful feedback on my project. I also thank Dr. William Merrick for giving me eIF4A and for reading the draft of this thesis. I thank Dr. Vernon Anderson for giving me RNA footing printing agent peroxynitrite and for his insight about enzyme kinetics. Many thanks to my advisor Dr. Eckhard Jankowsky. Without his continued support, it would have been impossible to finish this thesis. I am grateful for many insightful discussions with him, which were crucial to move the project forward. I also learned from him how to communicate scientific ideas effectively, including oral presentations and paper writing. Most importantly, the systematic way to investigate a problem I learned will continue to help me in my future career. I would like to thank Maggie Fairman for her initial introduction to the techniques for studying RNA helicases. I am in debt to her for letting me use her precious NPH-II preparation. I also thank her for explaining many crystal structures to me. I thank Heath Bowers for his initial studies on Ded1 and for his contribution to the idea of RNA structure conversion. His good ideas and insight about the helicase mechanism have been important to the progress of my project. I thank Dr. Nicholas Kay for helping me improve my proposal for PhD qualification and the introduction of this thesis. I want to thank Liu Fei for her technical support to plot the Figure 6.7 in this thesis. I thank Wen Wang for her optimization of the Ded1 purification and for helping construct Ded1 truncation mutants. I thank Dr. Mark Del Campo from Dr. Alan Lambowitz’s lab for producing purified Mss116. Finally, I would like to thank my parents and my wife for their unconditional love and support. I thank my daughter for highlighting my many days in Cleveland.

7 List of Abbreviations

ADP adenosine diphosphate AMPPNP adenosine 5' (beta, gamma-imido) triphosphate ATP adenosine triphosphate bp base pair BSA bovine serum albumin Ded1 defines essential domain 1 DTT dithiothreitol EDTA (ethylenedinitrilo)-tetra acetic acid eIF4A eukaryotic initiation factor 4A EJC HEPES N-(2-hydroxyethyl)-piperazine-N'-2-ethanesulfonic acid MOPS 3-morpholinopropanesulfonic acid NP40 nonidet-P40 NPH-II nucleotide phosphohydrolase II nt nucleotide NTP nucleotide 5’-triphosphate PAGE polyacrylamide gel electrophoresis PEI polyethyleneimine RNP ribonucleoprotein SDS sodium dodecyl sulfate TLC thin layer chromatography TRAP the trp RNA-binding attenuation protein Tris tris(hydroxylmethyl)-aminomethane wt wild type

8 Mechanism of RNA Remodeling by DEAD-box Helicases

Abstract

by

QUANSHENG YANG

DEAD-box proteins remodel RNA duplexes and displace proteins from RNA in

an ATP-dependent fashion. To understand how DEAD-box proteins remodel duplex

RNA, I studied the mechanism of RNA remodeling by the DEAD-box protein Ded1 from

S.cerevisiae. I found that Ded1 promotes not only RNA unwinding but also strand

annealing. Ded1 establishes an ATP-dependent steady state between unwinding and

annealing, which allows the enzyme to modulate the balance between the two opposing

activities through ATP and ADP. By coordinating its unwinding and annealing activities,

Ded1 can convert RNA structures via two distinct pathways. One pathway depends on

ATP hydrolysis and represents a kinetically controlled steady state between the RNA structures that involves the complete disassembly of the RNA structures. This pathway allows the formation of less stable RNA structures from more stable ones and thus

facilitates the distribution of RNA structures against thermodynamic equilibrium values.

The other pathway is ATP independent and involves stabilization of a multipartite

intermediate by Ded1. These results provide a basic mechanistic framework for a protein-

assisted RNA conversion that illuminates the role of ATP hydrolysis and an unexpected

diversity of pathways. I also discovered that Ded1 unwinds duplex RNA with a

mechanism distinct from “traditional” DNA helicases and some viral RNA helicases.

These “traditional” helicases unwind duplexes by first loading them to one of the two

nucleic acid strands, mostly at a single stranded region, and then by translocating on this

9 strand in a unidirectional fashion, thereby removing complementary strand. Ded1 does not unwind duplexes by translocating on a single stranded. Instead, a single stranded

RNA loads Ded1 to the duplex region, where strand separation is initiated directly. I also showed that the DEAD-box proteins Ded1 and Mss116 from S.cerevisiae can initiate duplex unwinding from internal positions of the duplex region. This new type of helicase activity explains puzzling observations with many other DEAD-box proteins and may be the prototype for duplex-unwinding reactions in RNA .

10 Chapter 1: Structures and biochemical activities of RNA helicases

1.1 Biological functions of RNA helicases

Ribonucleic acids () carry out a multitude of essential biological functions

ranging from telomere maintenance to the regulation of gene expression through pre-

mRNA splicing, RNA export, and mRNA translation (20, 21). In a physiological

environment, the structures of RNA are not static but highly dynamic (23).The dynamics

of RNA molecules involve alterations in RNA secondary and tertiary structure and many

of these alternations are coupled to the hydrolysis of adenosine triphosphate (ATP) (28).

It was thus a critical finding when enzymes were discovered that could catalyze

ATP-dependent conformational changes in RNA (39-42). Initially, these enzymes were

identified through their ability to alter complex RNA structures (39). Subsequently, it was

shown that these proteins could also separate complementary RNA strands driven by

ATP-hydrolysis (40, 42). Because this activity was reminiscent of the ability of DNA

helicases to unwind DNA helicases, the enzymes were designated RNA helicases (49).

This name is still widely popular, despite the fact that some of these enzymes also disrupt

RNA-protein (59-62) and, as recently shown, protein-protein complexes (63). Some of

these “RNA helicases” function as ATP-controlled clamps that facilitate assembly of

protein complexes on RNA (74-78).

RNA helicases can be found in all forms of cellular life (80). RNA helicases are

the largest group of enzymes in eukaryotic RNA metabolism and comprise approximately

0.6 % of the genes of S.cerevisiae (81-83). RNA helicases are involved in most, if not all, aspects of RNA metabolism, including , pre-mRNA splicing, biogenesis, RNA export, translation initiation, RNA degradation, mitochondrial gene

11 expression, and RNA editing (83, 93-106). RNA helicases have also been implicated in

cell differentiation (107) and cancer (108-110). Moreover, many viruses encode one or

more helicases (111) and often hijack host helicases to aid their replication (34, 35, 111,

113-117) Recently, it has been shown that RNA helicases also function as cellular receptors for viral RNAs that activate production of type I interferon (118-120).

Although, in a physiological context, RNA helicases function in highly specific reactions (106), the helicase activity generally does not require specific RNA sequences

when assayed in vitro (106), with one exception of the DEAD-box helicase DbpA, which

needs a RNA hairpin in rRNA to stimulate its ATPase and unwinding activities (31, 125,

126). The specificity of RNA helicases might therefore be conferred by means other than

specific RNA sequences. Conceivably, the targeting of RNA helicases is aided by protein cofactors (18, 19, 72, 105). However, it has also been shown that mitochondrial RNA helicases Mss116 and Cyt-19 bind group I and II introns nonspecifically and yet promote the specific splicing reactions (9, 129). Therefore, the signals that enable these enzymes to recognize their cognate substrates among the vast number of potential substrates in cells remain mysterious.

1.2 Characteristic sequence of RNA helicases.

RNA helicases are mostly members of the helicase superfamily 2 (SF2), although

there are some RNA helicases among SF1 (106). RNA helicases of SF2 share at least at

least eight characteristic sequence motifs that are conserved from bacteria to humans

(Figure 1.1A) (106). SF2 RNA helicases can be clearly divided into three subfamilies, the

DEAD-box, DEAH, and DExH helicases, named after the motif II in a single-letter

amino acid code (28, 106). SF2 RNA helicases are thus also called DExH/D helicases.

12 These conserved motifs are important for biochemical functions of DExH/D

helicases. Motifs I, II, and VI are involved in NTP binding and hydrolysis; motifs Ia, Ib,

IV and V are required for the RNA binding; motifs III and V couple NTP binding and

hydrolysis to strand separation (75, 77, 130-132). In addition, a newly identified Q motif,

upstream from motif I, confers ATP specificity (133). Although this Q motif, named after

the invariable glutamine residue, can be found in other helicases, it is most highly

conserved in DEAD-box helicases (133).

1.3 Three-dimensional structure of DExH/D helicases

Structures of several DExH/D helicases have been determined by X-ray

crystallography over the past decade(75, 77, 131, 132, 134-141). The three-dimensional

structures of DExH/D RNA helicases are strikingly similar to each other and to those of

SF1 DNA helicases, despite the large sequence divergence between SF1 and SF2 helicases (142, 143). The helicase core with the conserved characteristic motifs folds into

two highly similar RecA-like domains connected by a flexible peptide linker region (143)

(Fig.1.1B). Crystallographic studies have also shown that RNA helicases adopt at least

two conformations. In the “closed” conformation, binding of NTP brings the two RecA-

like domains together. Upon NTP hydrolysis and NDP/Pi dissociation, the two domains

swing apart and helicases adopt an “open” conformation (144). Therefore, the cycle of

NTP binding and hydrolysis translates into conformational changes of helicases, which,

conceivably, may be important for the function of these enzymes. However, how exactly

RNA helicases couple ATP binding and hydrolysis to conformational work on RNA is

not understood.

13 Figure 1.1

A

B

Figure 1.1 Structure and sequence characteristic of RNA helicases. (A) Characteristic sequence motifs of DExH/D helicases. DEAD-box, DEAH, and DExH at the left represent three subgroups of DExH/D helicases. The name of the conserved motifs is labeled on the top. The letters in the squared box represent the consensus sequences of respective motifs in the single-letter amino acid code and x represents any amino acid residue. Red boxes, motifs involved in NTP binding and hydrolysis; blue, RNA binding; gold, coupling of ATP binding and hydrolysis to duplex unwinding; purple, conferring ATP specificity in the DEAD-box subgroup. Caps, >75% conserved within each DExH/D helicases subgroup; Small caps, >50% conserved. (B) Side (left) and front (right) views of the DEAD-box helicase VASA in complex with RNA and nonhydrolysable ATP analog AMPPNP. Bound RNA is colored in green. AMPPNP is shown in a ball- and-stick representation. Locations of conserved sequence motifs are labeled in different colors. The circle represents the eIF5B binding site of VASA. The figure is adapted from reference (131).

14 1.4 Biochemical activities of RNA helicases

1.4.1 Nucleotide specificity of DExH/D helicases

All conformational work by RNA helicases, such as rearrangement of RNA structures and disruption of RNA-protein and protein-protein interactions, requires the hydrolysis of

ATP or other NTPs. Hydrolysis of NTP by RNA helicases is typically stimulated by

RNA binding (145-147). Most, if not all, DEAD-box helicases studied to date have a strong preference for ATP and 2-deoxy-ATP (dATP) (148) (Table 1.1). Other typical

(d)NTPs, i.e., (d)GTP, (d)CTP, and UTP (or TTP), are not hydrolyzed by most DEAD- box proteins. It is thought that ATP specificity of DEAD-box helicases is conferred by the Q-motif (133). Structural studies reveal that a conserved glutamine in the Q motif specifically recognizes ATP through interactions with the amino group in position 6 and imido group at position 7 of the adenine base (77, 131, 149). Nevertheless, several

DEAD-box helicases have been shown to hydrolyze other (d)NTPs as well. First, it is reported that DDX3 can hydrolyze all (d)NTPs (35). However, the DDX3 preparation was denatured by 6 M urea during the purification procedure (26) and the ATPase activity of this DDX3 preparation is RNA-independent, in contrast to the ATPase activity of DDX3 purified without denaturation, where the enzyme requires RNA (34,

35). Therefore, the exceptional RNA independent non-adenosine specific NTPase activity of DDX3 might be the result of enzyme denaturation. Second, DDX42 has been shown to hydrolyze all NTPs. However, ATP is preferentially hydrolyzed over other NTPs. In addition, only the ATPase, but not other NTPase activities, is stimulated by RNA (4).

Third, XP54 can hydrolyze at least GTP and ATP to support duplex unwinding (112).

15 Table 1.1 NTPase activities of DExH/D helicases

NTPasea NTP dNTP Sub- Protein Species group A C G U dA dC dG dT References

AN3 X.laevis DEAD + - - - + - - - (5, 6)

ATDRH1 A.thaliana DEAD + - - - + - - - (8)

CrhC A.sp DEAD + - - - + nd nd nd (10)

CrhR S.sp DEAD + - - - + - - - (2)

DBP5 S.cerevisae DEAD + - - - nd nd nd nd (17-19)

DbpA E.coli DEAD + - - - + nd nd nd (29-31)

DDX3 Human DEAD + + + + + + + + (34, 35)

DDX42 Human DEAD + + + + + nd nd nd (4)

Ded1 S cerevisae DEAD + - - - + - - - (7, 45)

eIF4A Human DEAD + - - - + - - - (14, 50, 51)

Has1p S cerevisae DEAD + - - - + - - - (57)

p68 Human DEAD + - - - + nd nd nd (42, 58)

p82 Human DEAD + - - - + nd nd nd (64)

RH-IIb Human DEAD + - - - + - - - (70, 71)

XP54 X.laevis DEAD + nd + nd nd nd nd nd (70, 71)

PRP2 S cerevisae DEAH + + + + + + + + (79)

PRP16 S.cerevisae DEAH + + + + + + + + (54-56)

PRP22 S.cerevisae DEAH + + + + + + + + (65-67)

PRP43 S.cerevisae DEAH + + + + + + + + (72, 73)

Brr2 S.cerevisae DExH + - - - + - - - (89-91)

NS3 BVDVd DExH + + + + + + + + (1)

NS3 DVe DExH + + + + nd nd nd nd (92)

NS3 GBVBf DExH + + + + + + + + (3)

NS3c HCVg DExH + + + + + + + + (12-14)

NS3c HGVh DExH + + + + + + + + (15) c Maleless D.melanogaster DExH + + + + + + + + (16)

NPH-IIc VVi DExH + + + + + + + + (24-27)

PPV-CI PPVj DExH + + + + + + + + (32, 33)

RHAc Human DExH + + + + + + + + (37, 38)

NSP2 SFVk DExH + + + + + + + + (121, 122)

SUV3 S.cerevisae DExH + + + + + + + + (123, 124)

TMP-CI TMPVl DExH + + + + nd nd nd nd (47)

U5200kD Human DExH + + - - + + nd nd (127, 128) a. Sign + indicates that the corresponding helicase can hydrolyze this type of NTP; - no NTPase activity; nd, not determined; b. also known as Gu; c. DNA helicase activity has also been shown. d. Bovine viral diarrhea virus; e. Dengue virus type 2; f. GB virus B; g. Hepatitis C virus; h. Hepatitis G virus; i. Vaccinia virus; j. Plum pox virus; k. Semliki Forest virus; l. Tamarillo mosaic potyvirus.

16 However, the XP54 preparation might be contaminated with other NTPases as XP54 is

purified by washing heterogeneous protein complexes bound to Poly(A)+ RNA over the

range of 0.4 - 1 M NaCl. These data suggest that even these proteins may eventually be

specific for ATP.

In contrast to DEAD-box helicases, most DEAH and DExH helicases do not have

a Q-motif and can hydrolyze all NTPs and dNTPs with an equal or similar efficiency to

support helicase activity (Table 1.1). Notwithstanding, it has been shown that the DExH

helicase Brr2 can only use ATP and dATP to support splicing reactions (89-91).

However, it remains to be seen whether Brr2 can hydrolyze other NTPs as well.

Promiscuity for nucleotide cofactors is also observed for SF1 DNA helicases (150-152).

1.4.2 Polarity of RNA duplex unwinding by DExH/D helicases.

Unwinding of RNA duplexes by many RNA helicases requires a single stranded

RNA adjacent to the duplex region (153, 154). The strand with this single stranded RNA

is termed as the loading strand. To unwind duplexes, helicases are thought to first bind to

this loading strand and then unidirectionally translocate on this strand through the duplex,

which causes strand separation (155). Many helicases that have been mechanistically

studied exclusively unwind substrates containing a single stranded region in one strictly defined orientation, either 5’or 3’ relative to the duplex (153-155). Helicases translocating in the 5’ to 3’ direction are thought to require the single strand 5’ to the duplex, those translocating 3’ to 5’ are assumed to need the single strand 3’ to the duplex.

17 Table 1.2 Polarity of RNA duplex unwinding by DExH/D helicases.

Unwinding Polarity a Protein Species Sub-group 3'->5' a 5'->3' a Blunt a References (1) NS3 BVDVe DExH Y(25) N(25) N(29) (3) NS3 GBVBf DExH Y(25) N(26) N(25) (12-14) NS3d HCVg DExH Y(25) N(26) N(11) (15) NS3d HGVh DExH Y(66) N(23) N(66) (16) Malelessd D.melanogaster DExH Y(30) N(30) N(30) (24-27) NPH-IId VVi DExH Y(120) N(12) nd (32, 33) PPV-CI PPVj DExH Y(17) N(17) nd (37, 38) RHAd Human DExH Y(25) N(26) N(25) (43, 44) hSUV3d Human DExH N(24) Y(32) nd (47) TMP-CI TMPVk DExH Y(16) nd nd

PRP16 S.cerevisiae DEAH Y(18) Y(18) nd (54-56)

PRP22 S.cerevisiae DEAH Y(30) N(30) N(30) (65-67) PRP43 S.cerevisiae DEAH Y(30) Y(30) nd (72, 73)

DEAD- DbpA E.coli box Y(10) N(10) N(10) (29-31) DEAD- DDX25 R.norvegicus box Y(20) Y(20) nd (46) DEAD- DeaDb E.coli box Y(14) Y(14) N(14) (52, 53) DEAD- Ded1 S.cerevisiae box Y(19) Y(16) Y(16) (7, 45) DEAD- DP103 Human box nd Y(15) nd (68, 69) DEAD- eIF4A Human box Y(13) Y(13) Y(13) (14, 50, 51) DEAD- Has1p S.cerevisiae box Y(16) Y(16) nd (57) DEAD- p68 Human box Y(22) Y(22) nd (42, 58) DEAD- RHLE E. coli box Y(14) Y(14) Y(14) (52) DEAD- RH-II c Human box N(25) Y(26) nd (70, 71) DEAD- SRMB E. coli box Y(11) Y(11) N(11) (52, 87) DEAD- TIF1/2 S.cerevisiae box Y(10) nd nd (88) DEAD- XP54 X.laevis box nd Y(46) nd (112) a. 3’Æ5’ refers to substrates with a single stranded region at the 3’-end; 5’Æ3’, substrates with a single stranded region at the 5’-end; Blunt, blunt-end substrates. Y refers that the corresponding helicase can unwind this type of substrates; N, can not; the number in the parenthesis indicates the duplex length used to examine the unwinding polarity; nd, not determined; b. also known as CsdA. c. also known as Gu; d. DNA helicase activity has also been shown; e. Bovine viral diarrhea virus; f. GB virus B; g. Hepatitis C virus; h. Hepatitis G virus; i. Vaccinia virus; j. Plum pox virus; k. Tamarillo mosaic potyvirus.

18 DExH helicases studied to date are invariably unidirectional and strictly require

an overhang, either 5’ or 3’ with respect to the duplex region, to initiate unwinding

(Table 1.2). This observation is consistent with the idea that strand separation is the consequence of unidirectional translocation of the enzymes along the loading strand.

Consistent with this notion, lesions such as nicks and polyglycol modifications on the

phosphodiester backbone of the loading strand, but not of the complementary strand (i.e., top strand), impede the unwinding activity of NPH-II (156). However, HCV NS3 can overcome a short stretch of polyglycol modifications in both its loading and top strands without decreasing the unwinding rate constants (157). Nevertheless, a longer stretch of polyglycol modifications and nicks in the loading but not in the top strand prevents unwinding by HCV NS3, supporting the idea that HCV NS3 has to translocate on the loading strand for strand separation.

The polarity of DEAH helicases is not clear (Table 1.2). While Prp22 has a strict requirement for a 3’ overhang and cannot unwind substrates with a 5’ overhang or blunt-

end substrates (67), Prp16 and Prp43 can unwind substrates regardless of the polarity of the overhang (55, 72). However, Prp16 unwinds substrates with a 3’ overhang more efficiently than those with a 5’ overhang (55), whereas Prp43 prefers substrates with a 5’

overhang over a 3’ overhang (72). Unwinding of blunt-end substrates by Prp16 and

Prp43, has not yet been reported and the unwinding activity for substrates with a “wrong”

overhang might reflect unwinding of blunt-end substrates.

Most DEAD-box helicases can unwind substrates regardless of the polarity of the

overhang (Table 1.2). Several enzymes have also been shown to unwind blunt-end substrates (Table 1.2) (51). Nevertheless, many DEAD-box helicases have poor

19 unwinding activity on blunt-end substrates and the presence of the overhang, regardless of its polarity, enhances unwinding rate constants (52, 53). While these results raised questions about unidirectional helicase translocation on the loading strand as a universal mode for duplex unwinding by all helicases(130, 153), it has remained unclear how strand separation can occur otherwise.

1.6 Processivity of DExH/D helicases

Helicases are thought to translocate unidireactionally in a stepwise manner. Step size is defined by how many nucleotides a helicase moves during one ATP hydrolysis cycle. An important characteristics of a helicase is its processivity, which is the probability of a helicase moving one step forward versus falling off the nucleic acid strand (154). A highly processive helicase can make multiple steps, even several hundreds, before falling off. Nonprocessive helicases can only make a few or just one step before falling off duplex RNA (148).

Initial studies on the prototypical DEAD-box helicase eIF4A suggested limited or absent processivity (158). In contrast, the DExH helicases NPH-II from vaccinia virus and HCV NS3(25, 159, 160) were later shown to be processive. Nonetheless, the approximately 40 DEAD-box and DEAH helicases studied so far appear more similar to eIF4A and show little or no processivity (130, 148) (Table 1.3). However, one should be cautious to judge the processivity based on the duplex length a helicase can unwind. A

20 Table 1.3 Unwinding capability of DExH/D helicases.

Sub- Protein Species group Duplex lengtha References DEAD- (5, 6) AN3 X.laevis box 10 DEAD- (8) ATDRH1 A.thaliana box 10 DEAD- (10) CrhC A.sp box 14

DEAD- (2) CrhR S.sp box 25 DEAD- (17-19) DBP5 S.cerevisiae box 10 DEAD- b (22) DBP9 S.cerevisiae box 17 DEAD-

DbpA E.coli box 12 (29-31) DEAD- DDX1 Human box 10 (36)

DEAD- DDX3 Human box 14 (34, 35) DEAD- DDX25 R.norvegicus box 20 (46) DEAD- c DeaD E.coli box 29 (48) DEAD-

Ded1 S.cerevisiae box 19 (52, 53) DEAD- DP103 Human box 15 (7, 45) DEAD- eIF4A Human box 13 (68, 69) DEAD- Has1p S.cerevisiae box 16 (14, 50, 51) DEAD- p68 Human box 162 (57) DEAD-

p72 Human box 25 (42, 58) DEAD- p82 Human box 25 (112) DEAD- RHLB E.coli box 11 (84) DEAD- RHLE E.coli box 14 (52)

DEAD- RH-II d Human box 26 (70, 71) DEAD-

SRMB E.coli box 11 (85) DEAD- TIF1/2 S.cerevisiae box 10 (86) DEAD- VASA D.melanogaster box 14 (52, 87) DEAD- XP54 X.laevis box 46 (88)

PRP16 S.cerevisiae DEAH 18 (54-56)

PRP22 S.cerevisiae DEAH 30 (65-67)

PRP43 S.cerevisiae DEAH 30 (72, 73)

NS3 BVDVe DExH 25 (1) NS3 DVf DExH 29 (92) NS3 GBVBg DExH 29 (3)

21

a. The longest duplex that has been shown to be unwound by the helicase; b. DNA helicase activity has also been shown; c. also known as CsdA; d. also known as Gu; e. Bovine viral diarrhea virus; f. Dengue virus type 2; g. GB virus B; h. Hepatitis C virus; i. Hepatitis G virus; j. Vaccinia virus; k. Plum pox virus; l. Semliki Forest virus; m. Tamarillo mosaic potyvirus.

22 helicase with pronounced annealing activity can catalyze strand exchange reactions in the

presence of a large excess of unlabeled homologous strand, which normally leads to

unwinding of much longer duplexes than without such trapping strand (cf. Chapter 4, and

(11)). As most unwinding reactions in the literature are conducted with a large excess of trapping strands, most unwinding data report large, artificially amplified unwinding capacities of RNA helicases

It should also be noted that exclusive RNA duplex structures exist mainly during viral replication and transcription. Contiguous RNA helices exceeding two turns are not normally seen in eukaryotic cells. Consequently, there appears to be little need for highly processive RNA helicases (28, 106). Nevertheless, it remains to be conclusively shown whether processivity is a feature characteristic of only the viral DExH helicases such as

NPH-II and HCV NS3.

1.7 Disruption of RNA-protein and protein-protein interactions by DExH/D

helicases.

In addition to their ability to unwind nucleic acid duplexes, several RNA helicases

have been directly shown to remodel RNA–protein complexes. The DExH helicase NPH-

II can displace protein complexes from both structured and unstructured RNA. Therefore,

the protein displacement is not necessarily coupled to duplex unwinding (61). Notably,

NPH-II can displace not only exon-junction complex (EJC) with a short binding site (8-9

nt), which is deposited on mRNA as a consequence of pre-mRNA splicing, but also the

trp RNA-binding attenuation protein (TRAP) complex with a much longer binding site

(53 nt) from single stranded RNAs (59, 61, 161). The dissociation of the TRAP complex

23 can be accelerated by more than three orders of magnitude by NPH-II compared with the

dissociation rate constant without NPH-II. The disruption of the TRAP complex requires

the presence of a single strand overhang, which is required by NPH-II for initiating

translocation. It is thus unlikely that NPH-II displaces the TRAP complex by acting

directly on the protein. Rather, these observations support the proposal that NPH-II

translocates on the ssRNA and that this translocation activity pushes the TRAP-complex

off the ssRNA (162).

However, translocation of DEAD-box helicases on a single stranded RNA may

not be required for protein displacement by nonprocessive DEAD-box helicase Ded1.

Ded1 can displace the EJC but not the TRAP complex, despite the fact that the EJC binds

more tightly on RNA than the TRAP complex (161). Conceivably, Ded1 can translocate on RNA a few steps to dislodge EJC but not processively enough to disrupt the RNA-

TRAP complex. Indeed, as for NPH-II, the EJC displacement by Ded1 also requires the

presence of a single stranded RNA, supporting the notion that Ded1 might also

translocate on the ssRNA. However, Ded1 does not use the overhang for initiating

translocation but for loading (Chapter 5). Therefore, it remains possible that Ded1

displaces EJC by directly acting on RNA-protein interactions without translocating on

RNA.

Recently, it has been reported that the DEAD-box helicase p68, when a tyrosine

residue at the C-terminus is phosphorylated, can displace Axin from β-catenin in an ATP-

dependent manner (63). The disruption of protein-protein interactions is likely not to

involve the translocation of p68 on ssRNA. Instead, these data indicate that p68 disrupts protein-protein interactions by acting directly on protein complexes, consistent with the

24 idea that Ded1 might not translocate on a single stranded RNA in order to displace the

EJC.

Collectively, these observations suggest that at least two different helicase modes among DExH/D helicases exists, unidirectional translocation on RNA and direct action on RNA-protein and protein-protein interactions without translocation. Moreover, these observations also expand the functional repertoire of DExH/D helicases and indicate that

DExH/D helicases might participate in biological processes not involving RNA.

1.8 Strand annealing activities of DExH/D helicases

Although the exact physiological targets for most DExH/D proteins are not well defined, these enzymes have been primarily associated with processes that involve the disassembly of RNA-RNA, RNA-protein, and even protein-protein interactions (163).

However, several DEAD-box helicases (Table 1.4) have recently been found to promote the formation of RNA duplexes in vitro, in addition to RNA unwinding (4, 11, 148, 164-

168). The annealing activity can be modulated by ATP and ADP. CrhR displays the annealing activity only in presence of ATP, whereas DDX42 requires ADP for the annealing activity. However, Ded1, Cyt-19, and Mss116 can promote strand annealing with and without ATP (unpublished data and ref. (7, 9)). The bimolecular strand annealing reaction can be accelerated at least three orders of magnitude, approaching the physical limitation imposed by diffusion(7, 9). The DExH helicase NPH-II and DEAD- box helicase eIF4A also promote the strand annealing reaction 10-fold, but to the extent

25 Table 1.4 A selection of DEAD-box helicases containing profound annealing activity.

Protein Species %Argininea %Lysinea References CrhR S. sp 10.39% 7.14% (2) Cyt-19 N. Crassa 11.24% 3.37% unpublished DDX42 human 7.49% 7.20% (4) Ded1 S.cerevisiae 8.04% 3.57% (7) Mss116 S.cerevisiae 9.74% 6.67% (9) p68 Human 7.78% 3.39% (11) p72 Human 7.34% 2.75% (11) SrmB E.coli 10.68% 19.42% unpublished a. The percentage of arginine and lysine at the C-terminus of the respective helicase immediately after the motif VI.

26 seen for the general RNA-binding protein U1A (discussed in Chapter 3). Therefore, the

profound annealing activity is not a general feature of DExH/D helicases but more likely a specific feature for a select number of DEAD-box helicases. Indeed, the DEAD-box helicases that contain the annealing activity share a region rich in positively charged amino acid residues at the C-terminus (Table 1.4). It has been shown that the positively charged peptide can promote the strand annealing by neutralizing the negatively charged phosphodiester backbone of nucleic acids (169, 170).

Proteins, which promote two processes that affect RNA secondary structure in

opposite directions, are potentially attractive points for regulating RNA structure

formation in complex biological systems. However, it is unclear whether DEAD-box

proteins that catalyze both RNA unwinding and duplex formation, have the capacity to

differentially affect either activity. In fact, there is little mechanistic understanding how

these DEAD-box proteins coordinate RNA unwinding and annealing activities.

1.9 Ded1 as a model system to study the molecular mechanism of RNA remodeling

by DEAD-box helicases

Data accumulated over the last decade suggest that DEAD-box helicases differ

from DExH helicases in several aspects: 1) DEAD-box helicases unwind substrates

regardless of the polarity of the overhang, whereas DExH helicases only unwind

substrates with an overhang in a strictly defined orientation, either 5’ or 3’ with respect to

the duplex region; 2) DEAD-box helicases disrupt RNA-protein and protein-protein

interaction without translocation, whereas some DExH helicases disrupt RNA-protein

interactions by translocating processively on ssRNA. These observations are inconsistent

27 with unidirectional translocation on ssRNA as universal remode for RNA/protein modeling by all RNA helicases. However, it remains to be shown how RNA/protein remodeling can occur otherwise. To gain insight how DEAD-box helicases remodel duplex RNA, I have studied in vitro RNA remodeling by the DEAD-box helicase Ded1 from S. cerevisiae.

Ded1 (definition of essential domain 1) (83, 171) features all of the conserved

sequence motifs characteristic of DEAD-box helicases, in addition to arginine-glycine-

aspargine rich C- and N-termini (7, 45). Ded1 is highly similar (72% identical) to another

DEAD-box helicase, Dbp1, in yeast (172). However, functions of Ded1 appear not

completely redundant with Dbp1, as deletion of Ded1 is lethal (83). The mRNA level of

Dbp1 is 10-fold lower than that of Ded1 (172, 173). This low expression level might

explain that the loss of function due to deletion of Ded1 cannot be completely

compensated by endogenously expressed Dbp1 but can be rescued by Dbp1

overexpression (172). Unlike Ded1, Dbp1 can be deleted without an obvious phenotype

(83). Ded1 has homologues and orthologues in the higher organisms (174), including

Belle in D. melanogaster (175), An3 in X. laevis (6), PL10 in mice (176), and DDX3 and

DBY in human (177, 178). Ded1 is also closely related vasa-like proteins such as Dbp2 in S. cerevisiae, VASA in X. laevis, and DDX4 in human (174, 179). It has been shown that DDX3 from human, as well as Belle from D. melanogaster, can rescue the Ded1 null mutation in yeast but not vice versa (115, 180).

Ded1 has been implicated in multiple cellular processes, including translation

initiation (181-183), pre-mRNA splicing (184, 185), and ribosome biogenesis (186).

Ded1 was first isolated by Struhl in 1985 as an essential gene in yeast (171). Later, Ded1

28 was also isolated from a screen for suppressing mutations of prp8, a protein required for pre-mRNA splicing (184). In accordance with its role in pre-mRNA splicing, Ded1 was isolated as a component of the yeast spliceosomal penta-snRNP (185). It was also shown

that Ded1 genetically interacted with a subunit of RNA polymerase III (187), and was

isolated from pre-ribosomal particles (186). However, it is unclear which roles (if any)

Ded1 plays in transcription or ribosome biogenesis. Apart from its nuclear roles, Ded1 is

required for translation initiation in the cytoplasm. Depletion of Ded1 in an in vitro yeast

translation system abolishes translation and this defect can be rescued by addition of

recombinant Ded1 (183). In addition, the fraction of monosomes in the ribosome profile

increases when yeast harboring a cold sensitive Ded1 mutation was shifted to the

nonpermissive temperature, suggesting a defect in translation initiation. However, it is

not clear which exact role Ded1 plays in translation initiation. Ded1 was also isolated

from screens for the PAS-domain kinase psk1 psk2 mutation (188) in S. cerevisiae and

sterility mutants (189) and Cdc25 overexpression (190) in S. pombe. Ded1 in S. pombe

has been linked to the cell cycle by regulating the essential cell cycle proteins including

cyclin-dependent kinase cdc2, cig2, and Cdc13 (190). Interestingly, Ded1 overexpression also enhances gene silencing by antisense RNAs (191). Although it is exciting to think that Ded1 might have many roles in these cellular processes, the observations might also reflect translational control by Ded1 of proteins involved in all of these processes(174).

Ded1 has been shown previously to display RNA-dependent ATPase activity and ATP-

dependent helicase activity in vitro (45).

To test whether the RNA remodeling mechanism observed for Ded1 is a general

feature of DEAD-box helicases, I have also used Mss116 from S. cerevisiae, which is

29 phylogenetically distant from Ded1within the DEAD-box helicase family. Mss116 also has RNA-dependent ATPase activity and ATP-dependent unwinding activity in vitro (9).

As for Ded1, Mss116 contains a C-terminus rich in positively charged residues and displays pronounced strand-annealing activity in vitro (9). Although Mss116 is not essential for yeast growth (83), it is required for splicing of mitochondrial group I and II introns in vivo and in vitro (9, 192). Cyt-19 from Neurospora crassa is an orthologue of

Mss116. Consistent with this notion, the functions of Mss116 in yeast can be substituted by Cyt-19 (192).

30 Chapter 2: Initial characterization of Ded1

Deciphering the mechanism by which Ded1 unwinds and remodels RNAs

requires first the establishment of the reaction conditions that allow the generation of

quantitative data for unwinding reactions in a reproducible and efficient manner. These

issues are discussed in the following sections.

2.1 Results:

2.1.1 Expression and purification of Ded1.

Bacterial cells with a pET22b plasmid containing the Ded1 gene (45) were grown

in LB media. When the absorbance at 600 nm (A600) reached to ~0.6, the expression of

Ded1 was induced by addition of 0.5 mM IPTG (final concentration) and the induction

was continued overnight at 28 ºC (45, 61). When expressed at higher temperature such as

37 ºC, most Ded1 was found in the insoluble fraction after lysis of bacterial cells.

Ded1 was purified by absorption on Ni-Agarose beads (Qiagen) in buffer A

(material and method) and subsequent elution with 250 mM imidazole. Ded1 was then

further purified by absorption onto Phosphocellulose (P-11, Whatman) and elution with

300 mM NaCl in buffer B (material and method). The purity of Ded1 was greater than 95

%, as judged by the Coomassie staining of purified Ded1 on the sodium dodecyl sulfate - polyacrylamide gel electrophoresis (SDS-PAGE) gels. The protein concentration of Ded1

preparations was determined by the Bradford method with bovine serum albumin (BSA)

as standard. Attempts to prepare Ded1 with concentrations higher than 7 μM were

unsuccessful in buffers including Tris-HCl (pH 8.0), MOPS (pH 6.5), HEPES (pH 7.2),

31 and phosphate buffer (pH 6.5 or 8.0), as Ded1 was prone to precipitation. Ded1 displayed

best solubility in Tris-HCl buffer (pH 8.0) among the buffers I examined. Ded1 was

aliquoted and stored at -80 °C. The activity of Ded1 did not decrease appreciably in

storage at -80 °C for 12 months.

2.1.2 Ded1 has RNA-dependent ATPase activity and ATP-dependent unwinding

activity.

To assess the functional properties of Ded1, the ATPase activity of Ded1 was

first examined. As reported by others (45), ATP was hydrolyzed by Ded1 in the absence

of RNA (Fig.2.1 B), with a slow rate of 3.1 min−1 with 100 nM Ded1 and 0.5 mM ATP at room temperature. Ded1’s ATPase activity was stimulated by one µM RNA (The ATPase

activity of Ded1 is saturated by RNA at this concentration, data not shown) (Fig.2.1

A,B), with its rate increased 11.1-fold compared with the reaction without RNA, to 34.8 min−1. Next, the unwinding activity of Ded1 was investigated. The 13 bp duplex RNA with a 25 nt ssRNA at the 3’-end (Table 2.1, substrate C) was unwound readily by Ded1 in the presence of ATP, with a rate constant of 2.82 min−1(Fig.2.1 C,D). The observed

unwinding rate constants varied about 20% among different Ded1 preparations. Ded1 was not able to unwind 13 bp duplex RNA in the absence of ATP after incubation of

Ded1 with duplex RNA for 5 min (Fig 2.1C, lane zero). These data demonstrate that

Ded1 has ATP-dependent helicase activity and the unwinding rate constants can be

generated reproducibly even when different Ded1 preparations are used.

32 Figure 2.1

AB 1.0 Pi 0.8 0.6 0.4 ATP 0.2

Fraction Phosphate Fraction 0.0 0 0.1 40 0 10 20 30 40 Time [min] Time [min] CD 1.0 * 0.8 0.6

* 0.4 0.2 0 2

Fraction Unwound Fraction 0.0 Time [min] 0.0 0.5 1.0 1.5 2.0 Time [min]

Figure 2.1 Ded1 has RNA-dependent ATPase activity and ATP-dependent unwinding activity. (A) Representative PEI TLC plate for the ATPase reaction with 100 nM Ded1 and 0.5 mM ATP plus a trace amount of [32P -γ-ATP] in the presence of 1 μM 41 nt ssRNA (Table 2.1, substrate A). Reactions were performed at room temperature. Mobility of [32P -γ-ATP] and the inorganic [32P] on the TLC plate are indicated on the left. Zero represents the ATP prior to addition to the ATPase reaction mixture. Aliquots were taken from 0.1 to 40 min. (B) Time courses of the ATPase reactions without RNA (●) or with 41 nt (○) RNA. RNA is the same as in panel A. Data points represent the average values from at least two independent reactions. The lines represent a linear trend. (C) Representative PAGE of unwinding reactions with 100 nM Ded1 and 2 mM ATP for the 13 bp RNA duplex with a 25 nt ssRNA at the 3’-end (Table 2.1, substrate C). Reactions were performed at room temperature. Cartoons on the left represent mobility of duplex and single strand. Asterisks represent the radiolabels. Zero represents the aliquot taken after incubation of duplex RNA with Ded1 for 5 min but prior to ATP addition. After addition of ATP, aliquots were removed from 5 s to 2 min. (D) Time course of the unwinding reaction shown in panel C. Data were fitted to the integrated first order rate law (Materials and Methods, eq III), yielding reaction amplitude of A (unw) −1 = 0.996 ± 0.063 and kobs = 2.82 ± 0.44 min .

33 Table 2.1 Substrates and their sequences

Substrate Sequence

A 5’-UUAGUACGUCCCAGACAGCAUUGUACCCAGAGUCUGUACGG-3’

5’-AGCACCGUAAAGA-3’ B ||||||||||||| 3’-UCGUGGCAUUUCU-5’

5’-AGCACCGUAAAGA-3’ C ||||||||||||| 3’- AAAACAAAACAAAACAAAACAAAAUUCGUGGCAUUUCU-5’

34 2.1.3 The effect of pH of the reaction buffer on the unwinding activity of Ded1.

To find the optimal conditions under which the unwinding rate constants of Ded1

could be accurately measured by manual pipetting and without significant self-cleavage

of the RNA, the effect of pH on the unwinding activity of Ded1 for a 13 bp RNA substrate with a 25 nt ssRNA region at the 3’-end (Table 2.1, C) was investigated. The unwinding activity of Ded1 was not optimal at pH 7.0, the physiological pH value in yeast (Fig2.2A). Lowering the pH value from 7 to 6.5 decreased the unwinding activity of Ded1 further (Fig2.2A). The unwinding activity of Ded1 at pH 8.0 was higher than at

pH 7.0 and pH 6.5 (Fig2.2A).

Ded1 was also able to unwind 13 bp blunt-end RNA duplex (Table 2.1, B) but this reaction proceeded slower than that for 13 bp duplex with a 25 nt overhang at the 3’-

end. The unwinding activity of Ded1 for the blunt-end duplex was also higher at pH 8.0

than pH 7.0 and pH 6.5 (Fig 2.3 A). Although the unwinding activity of Ded1 at the

greater pH value might be higher than at pH 8.0, I did not examine these conditions

because pH greater than 8.0 adversely affects the stability of RNA in aqueous solution,

especially in the presence of Mg2+ ion (193).

2.1.4 The unwinding activity of Ded1 is inhibited by increasing salt concentrations.

Next, how the salt concentration affected the unwinding rate constants of Ded1 at

pH 8.0 for the 13 bp duplex with a 25 nt ssRNA overhang (Table 2.1, substrate C) was

investigated. Increasing NaCl concentrations repressed the unwinding activity of Ded1.

The rate constant decreased ~2600-fold for an increase of NaCl concentrations from 20

mM to 220 mM. (Fig 2.2 B). KCl inhibited Ded1’s unwinding

35 Figure 2.2

A

] B -1 100

10

1

0.1

0.01

Observed Unwinding Rate Constant [min 6.0 7.0 8.0 0 50 100 150 200 pH NaCl [mM] C D ] -1 100

10

1

0.1

0.01

RateConstant [min 0 50 100 150 200 Observed Unwinding 02468 KCl [mM] MgCl2 [mM] Figure 2.2 Effects of the pH and salt concentration on the unwinding activity of Ded1 for 13 bp duplex

RNA containing a 25 nt ssRNA at the 3’-end (Table 2.1, substrate C) with 100 nM Ded1 and 2 mM

ATP/MgCl2 at room temperature. (A) Dependence of unwinding rate constants on pH. The line through

the data points represents a linear trend. (B) Dependence of unwinding rate constants on the NaCl

concentrations. (C) Dependence of unwinding rate constants on the KCl concentrations. (D) Dependence of

unwinding rate constants on the MgCl2 concentrations. The MgCl2 concentrations are the free MgCl2 in

addition to the 2 mM MgCl2 that accompanies ATP.

36 Figure 2.3

] A B -1 1

0.1

0.01

0.001 Observed Unwinding

Rate Constant [min 6.0 7.0 8.0 020406080 100

pH NaCl [mM] C D ]

-1 1

0.1

0.01

0.001

RateConstant [min 020406080 100 02468 Observed Unwinding KCl [mM] MgCl2 [mM]

Figure 2.3 Effects of the pH and salt concentration on the unwinding activity of Ded1 for 13 bp blunt-end duplex. (A) Dependence of unwinding rate constants on pH. (B) Dependence of unwinding rate constants on the NaCl concentrations. (C) Dependence of unwinding rate constants on the KCl concentrations. (D)

Dependence of unwinding rate constants on the MgCl2 concentrations.

37 activity to a similar extent (Fig 2.2 C). The divalent salt MgCl2 was almost twice as potent as the monovalent salts NaCl and KCl for the inhibition of Ded1’s unwinding activity (Fig 2.3 D). Increasing salt concentrations also suppressed the unwinding for the blunt-end substrates. NaCl is as efficient as KCl in inhibiting the unwinding rate constant for the blunt-end substrate within experimental error, with its rate constants reduced ~60 fold for an increased of salt from 20 mM to 80 mM (Fig 2.3B,C). MgCl2 was more

effective than NaCl and KCl in inhibiting the unwinding activity of the blunt-end duplex

(Fig 2.3 D).

These data suggest that increasing ionic strength might suppress the unwinding activity of Ded1. The higher efficiency of MgCl2 than NaCl and KCl in inhibiting Ded1’s

unwinding activity might reflect the higher capability of the divalent salts to raise the

ionic strength than the monovalent salts. It has been shown that DEAD-box helicases

associate with duplex RNA mainly through the electrostatic interactions (75, 77, 131).

Therefore, at higher ionic strength, binding of Ded1 with duplex RNA might be impaired,

which in turn results in inhibition of the unwinding activity.

To perform the functional studies on Ded1, the following reaction conditions, 50

mM NaCl, 0.5 mM MgCl2 and pH 8.0, were chosen.

38 2.2 Discussion

In this Chapter, I have shown that Ded1 has RNA-dependent ATPase activity and

ATP-dependent unwinding activity. The unwinding activity of Ded1 is inhibited by

increasing salt concentrations.

The helicase core of DEAD-box helicases folds into two highly similar RecA-like

domains connected by an extended peptide linker region in the absence of RNA and ATP

(137). In the presence of RNA and ATP, two RecA-like domains swing closer (75, 77,

131). The RNA stimulated ATPase activity observed for the DEAD-box helicases Ded1,

CsdA (52), eIF4A (50), Has1p (57), RhlE (52), and SrmB (52, 87) and the DEAH helicases like Prp22 (67) and Prp43 (72) suggest that RNA binding to these helicases promote orientation of two RecA-like helicase domains that is required for ATP binding and/or hydrolysis.

However, some DEAD-box helicases such as Dbp8 and Dbp9 do not require

RNA to stimulate their ATPase activities (22, 194). In fact, the ATPase activity of Dbp9

is inhibited by RNA (22). The lack of the ATPase stimulation by RNA cannot be

explained by the possible contamination of RNA in the enzymes after the protein

purification, as their ATPase activities after the RNase treatment remain independent of

RNA. DNA can stimulate the ATPase activity of Dbp9, consistent with the DNA helicase

activity it contains (22). These data indicate that some DEAD-box helicases are not RNA

helicases but DNA helicases.

The ATPase assays of helicases are often carried out with 5 mM Mg2+ and 100

mM NaCl or KCl. The ATPase activity is readily detectable under these conditions.

However, high salt concentrations are detrimental for the unwinding activity of most

39 RNA helicases, including Ded1, eIF4A(158), and NS3 (195). NS3 is particularly sensitive to Mg2+ concentrations. Small amounts of excess Mg2+ other than the Mg2+

bound to ATP results in significant loss of the unwinding activity of NS3. Presumably,

increasing the divalent ion concentrations weakens the interaction between the enzymes

and their RNA substrates, which, in turn, reduces their ATPase and unwinding activities.

Indeed, RNA helicases interact with their RNA substrates mainly through electrostatic

interactions (77, 131, 132, 196) and low ionic strength reaction conditions can therefore

promote the binding of the enzymes to their RNA substrates(197). Alternatively,

increasing ion concentrations stabilizes RNA duplexes by neutralizing the negatively

charged phosphate backbone of the RNA strands. It has been shown that the unwinding

ability of nonprocessive DEAD-box helicases including eIF4A and Ded1 decreases

dramatically with increasing stability of RNA duplexes (7, 51).

40 Chapter 3: ATP and ADP-dependent modulation of RNA unwinding and strand annealing activities by the DEAD-box helicase Ded1

The unwinding rate constants observed in the previous Chapter most likely

describe a multi-step reaction including the forward unwinding reaction and possibly its reversal strand-annealing reaction, i.e., formation of duplex RNA from its respective

RNA strands. To decipher the unwinding mechanism, the effect of the annealing reaction

on the observed unwinding rate constants requires a careful quantification in order to

interpret the observed rate constants in a meaningful manner. In this Chapter, I describe

the studies on the strand annealing activity of Ded1.

41 3.1 Results

3.1.1 Ded1 catalyzes both unwinding and annealing of RNA duplexes.

To analyze the unwinding activity of Ded1, I first tested the ability of the enzyme to unwind an RNA duplex substrate containing a 25-nucleotide overhang

3’ to a 16 basepair duplex region (Table 3.1A). Ded1 readily separated the two strands in the presence of ATP (Fig.3.1A). Unwinding was only observed with ATP or dATP

(Fig.3.1A, B). Virtually no helicase activity was detected with other nucleoside triphosphates and no significant strand separation occurred in the presence of the ATP analog AMPPNP or in the presence of ADP (Fig.3.1B). These results show that the RNA helicase activity of Ded1 is strictly dependent on not only the binding of adenosine triphosphates but also hydrolysis.

Ded1 also significantly accelerated the formation of an RNA duplex from the single strands of the above substrate (Fig.3.1C,D). This annealing activity did not require ATP

(Fig.3.1C). To test whether the strand annealing activity was specific for Ded1, the extent to which duplex formation was promoted by two other DExH/D enzymes, the DExH helicase NPH-II (24, 25, 198) and the DEAD-box helicase eIF4A, was measured (51,

158) (Fig.3.2). Both proteins increased the observed annealing rate constant by approximately one order of magnitude over the basal rate constant (Fig.3.2A). Ded1, in contrast, increased the annealing rate constant by more than three orders of magnitude

(Fig.3.2A). These findings suggest that the pronounced catalysis of duplex formation is specific for Ded1 and not a general feature of DExH/D proteins.

The capacity to catalyze both duplex unwinding and strand annealing in vitro had been observed for four other DEAD-box proteins, human p68 and p72, yeast Mss116,

42 Table 3.1 Substrates and their sequences

Substrate Sequence

S[16] 5’-AGCACCGUAAAGACGC-3’ |||||||||||||||| 3’- AAAACAAAACAAAACAAAACAAAAUUCGUGGCAUUUCUGCG-5’

5’-AAAACAAAACAAAACAAAACAAAAU F[16] AGCACCGUAAAGACGC-3’ |||||||||||||||| 3’- AAAACAAAACAAAACAAAACAAAAUUCGUGGCAUUUCUGCG-5’

5’-AGCACCGUAAAGACGCAGC-3’ S[19] ||||||||||||||||||| 3’- AAAACAAAACAAAACAAAACAAAAUUCGUGGCAUUUCUGCGUCG-5’

43 Figure 3.1

A B * *

* * 0 5 95° A U G C A T G C AD Time [min] C NTP dNTP P C D

* *

* * 0 3 C 0 3 Time [min]0 Time [min] 0 Figure 3.1. RNA unwinding and strand annealing activities of Ded1. (A) Representative PAGE of an unwinding time course. Mobilities of duplex and single stranded RNAs are indicated by the cartoons on the left. The arrow emphasizes that the reaction was started with duplex RNA. The zero timepoint represents the reaction before ATP addition. Aliquotes were removed between 5 s and 5 min. (B) Unwinding reactions with 1 mM various NTPs (left panel), AMPPNP and ADP (right panel). Reactions were performed as described in the methods section except that ATP-MgCl2 was replaced with equimolar mixtures of MgCl2 and NTPs, dNTPs, AMPPNP, or ADP, as indicated underneath the gels. Reactions were incubated for 30 min. (C) Representative PAGE of a strand annealing time course with Ded1. No ATP was present during the reaction. The arrow indicates that the reaction was started with single stranded RNA. The zero time point represents the reaction before Ded1 addition. Aliquots were removed from 0.5 min to 30 min. (D) Representative PAGE of a strand annealing time course without Ded1. Reactions were performed as in panel (C), except that Ded1 was omitted.

44 and CrhR from cyanobacteria (2, 9, 11). These proteins share with Ded1 a characteristic

RK-rich C-terminus, in addition to the commonly conserved regions of the DEAD-box

helicases (Fig.3.2B). To ascertain the importance of the C-terminus for the annealing

activity of Ded1, the C-terminal 83 amino acids from Ded1 were deleted. The mutant

protein Ded1ΔC was expressed and purified. The annealing activity of this altered peptide

(Fig.3.2A) was then measured. Ded1ΔC displayed an approximately 12-fold decreased rate constant of duplex formation, compared to wild type Ded1 (Fig.3.2A), whereas the unwinding activity was only reduced by a factor of ~ 1.2 (cf. Fig.3.3F). This result suggests that the C-terminus with the characteristic RK-rich region contributes to the annealing activity of Ded1. Yet, the ability to promote duplex formation does not exclusively reside within this protein segment.

3.1.2 Ded1 establishes an ATP-dependent steady state between RNA unwinding and strand annealing.

Having established the ability of Ded1 to catalyze the formation of RNA duplexes in the

absence of ATP, I next investigated whether Ded1 also promoted strand annealing in the

presence of ATP. With 0.1 mM ATP, Ded1 readily catalyzed duplex formation

(Fig.3.3A). However, the reaction did not proceed to completion, as observed without

ATP (Fig.3.1C). Instead, a distinct reaction amplitude was reached (Fig.3.3A,C). This

reaction amplitude was not caused by inactivation of the enzyme or ATP depletion during

the reaction, as Ded1 remained fully active for more than 60 min and consumed less than

1% of ATP under all reaction conditions (data not shown).

Unwinding time courses at 0.1 mM ATP also showed a clear reaction amplitude

45 Figure3.2

A

BSA control

NPH-II

eIF4A

DED1

DED1ΔC 105 106 107 108 109 1010

Annealing Rate Constant [M-1min-1]

B Motif VI p68 YPNSSEDYIHRIGRTARSTKTGTAYTFFTPNNIKQVSDLISVLREANQAINPKLLQLVED 477 p72 YPNSSEDYVHRIGRTARSTNKGTAYTFFTPGNLKQARELIKVLEEANQAINPKLMQLVDH 475 DED1 LPSDVDDYVHRIGRTGRAGNTGLATAFFNSENSNIVKGLHEILTEANQEVPSFLKDAMMS 535 CrhR LPDNAETYIHRIGRTGRAGKTGKAIALVEPIDRRLLRSIENRL-KQQIEVCTIPNRSQVE 380 Mss116 VPSELANYIHRIGRTARSGKEGSSVLFICKDELPFVRELEDAKNIVIAKQEKYEPSEEIK 512 p68 R-----GSGRSRGRGG------MKDDRRDRYSAGKRGG----FNTFRDRENYDRG-YS 519 p72 RGGGGGGGGRSRYRTTSSANNPNLMYQDECDRRLRGVKDGGRRDSASYRDRSETDRAGYA 535 DED1 AP-----GSRSNSRRGGFGRNNNRDYRKAGGASAGGWGSSRSRDNSFRGGSGWG------589 CrhR [381-448]KPVLRRGRNAGGGQNKSGGGYQGKPGKPRRSSGGRRPAYSDRQQ------492 Mss116 [513-617]RGNKNYNNRSQNRDYDDEPFRRSNNNRRSFSRSNDKNNYSSRNSNIY---- 664

Figure 3.2. Pronounced strand annealing activity is specific for Ded1. (A) Observed rate constants for strand annealing reactions (S[16]) in the presence of different proteins. Points on the scaled bars (note log ann scale) indicate the observed annealing rate constants (kobs ), which were calculated by fitting time courses of annealing reactions to eq. IV (Materials and Methods). Time courses for each protein given on the left were constructed from at least six time points. Each time point was determined in at least three independent experiments. All reactions were performed at identical salt and buffer conditions with identical RNA concentrations (Materials and Methods). Proteins were present in the reaction in the following concentrations: BSA: 1 μM, control: no protein added, NPH-II: 50 nM (saturating with respect to the RNA, (25), eIF4A: 1 μM (maximal possible concentration, (158), Ded1: 600 nM (saturating with respect to

the RNA, Materials and Methods), Ded1ΔC: 600 nM (saturating with respect to the RNA, data not shown).

Ded1ΔC lacked the C-terminal 83 amino acids (Materials and Methods). (B) Sequence alignment of the C- termini of DEAD-box proteins with annealing activity in vitro. Alignments were generated with CLUSTAL X (199). Numbers indicate amino acids counted from the N-terminus.

46 that was virtually identical to the amplitude of the annealing reaction at the same ATP

concentration (Fig.3.3 B,C). At 0.25 mM ATP, reaction amplitudes for the unwinding

and annealing process were virtually identical, too (Fig.3.4D). Notably, a larger fraction

of single stranded RNA was generated at 0.25 mM ATP than at 0.1 mM ATP (Fig.3.3D).

These observations show that Ded1 generated the same fraction of duplex and single

stranded RNAs at a given ATP concentration, regardless of whether the reaction was

initiated from single strands or from the duplex. Thus, observed amplitudes are not the

result of a stable dead-end complex between Ded1 and one (or both) of the RNA strands,

because then unwinding would have proceeded to completion, as the dead-end complex

would have “pulled” all RNA towards the single stranded products. Instead, the data were

consistent with Ded1 simultaneously annealing and unwinding the RNA until a steady

state between duplex and single stranded RNA was reached. Since the amplitude of a

steady state between two opposing processes is determined by the ratio of the two

reaction rates, greater reaction amplitude at a higher ATP concentration further suggested

that increasing ATP concentrations promoted the rate of the unwinding process, relative

to the annealing reaction. To confirm this assertion, time courses of unwinding reactions

over a larger range of ATP concentrations were measured (Fig.3.3E). Reaction amplitudes (fraction of single stranded RNA) increased with the ATP concentrations, further supporting the notion that in a steady state between duplex unwinding and formation, higher ATP concentrations accelerated the rate of unwinding, relative to the rate of annealing.

To further establish the ability of Ded1 to promote an ATP-dependent steady state

between unwinding and annealing, a central prediction of this mechanism was directly

47 Figure 3.3

A B annealing unwinding

* *

* *

0 30 0 30 Time [min] Time [min]

C D ATP = 0.1 mM ATP = 0.25 mM

1 1 0.8 annealing 0.8 annealing 0.6 0.6 0.4 0.4 Fraction

Single Strand Single 0.2 unwinding 0.2 unwinding

0 0 0 5 10 15 20 25 30 0 5 10 15 20 25 30 Time [min] Time [min] E F

wt DED1 DED1ΔC 1 1 0.8 0.8 0.6 0.6

0.4 0.4 Fraction Fraction

Single Strand 0.2 0.2 0 0 0 5 10 15 20 25 30 0 5 10 15 20 25 30 Time [min] Time [min]

Figure 3.3 Ded1-catalyzed ATP-dependent steady state between RNA unwinding and strand annealing. (A) Representative PAGE for a strand annealing reaction in the presence of Ded1 and 0.1 mM ATP. The RNAs were identical to those used in Figure 3.1. The arrow indicates that the reaction was started with single stranded RNA. Aliquots were removed from the reaction at the time indicated in panel (C). (B) Representative PAGE for an unwinding reaction in the presence of Ded1 and 0.1 mM ATP. The arrow indicates that the reaction was started with duplex RNA. Aliquots were removed at the time indicated in panel (A). (C) Time courses of strand annealing (○) and unwinding (●) reactions in the presence of Ded1 and 0.1 mM ATP. Data points represent the average of at least three independent reactions, error bars indicate one standard deviation. The line through the data points represents the best fit to the integrated

48 form of a homogenous first order rate law (eq. III, Materials and Methods). The reaction amplitude Aunw = 0.41 ± 0.02. The line through the annealing time course represents the best fit to the integrated form of a homogenous first order rate law (eq. IV, Materials and Methods). Aann = 0.43 ± 0.02. (D) Time courses of strand annealing (○) and unwinding (●) reactions in the presence of Ded1 and 0.25 mM ATP. Data were measured and rate constants and reaction amplitudes were determined as in panel (C). Aunw = 0.56 ± 0.05, Aann = 0.54 ± 0.02. (E) Time courses for unwinding reactions with wtDed1 at 0.1 mM ATP (●), 0.5 mM ATP (○) and 2 mM ATP (♦). Time courses were fitted to eq. III, yielding the following values: 0.1 mM unw -1 unw unw -1 unw ATP: kobs = 1.5 ± 0.1 min , A = 0.42 ± 0.02, 0.5 mM ATP: kobs = 3.0 ± 0.2 min , A = 0.73 ± unw -1 unw 0.02, 2 mM ATP: kobs = 3.7 ± 0.4 min , A = 0.90 ± 0.05. (F) Timecourses for Ded1ΔC catalyzed unwinding reactions at 0.1 mM ATP (●), 0.5 mM ATP (○) and 2 mM ATP (♦). Time courses were fitted to unw -1 unw eq. III, yielding the following values: 0.1 mM ATP: kobs = 0.6 ± 0.2 min , A = 0.90 ± 0.02, 0.5 mM unw -1 unw unw -1 unw ATP: kobs = 2.5 ± 0.3 min , A = 0.97 ± 0.03, 2 mM ATP: kobs = 2.8 ± 0.3 min , A = 0.97 ± 0.03.

49 tested: reaction amplitudes at a given ATP concentration result from a distinct ratio of the

rates for both unwinding and annealing. Conceivably, measuring the ATP-dependence of

reaction amplitudes with a variant of Ded1 that promoted both processes with a different

velocity ratio should yield different amplitudes, compared to reactions with wtDed1

under identical reaction conditions. Therefore, unwinding reactions with the C-terminal

Ded1 deletion mutant, Ded1ΔC, were performed. Compared to wt Ded1, the annealing

activity of Ded1ΔC was reduced by a factor of 12, whereas the unwinding activity was decreased only by a factor of 1.2 (Figs.3.2A, 3.3F). Thus, Ded1ΔC inherently favored

unwinding over annealing, compared to wtDed1, and higher amplitudes for unwinding

reactions with Ded1ΔC were anticipated. This expectation was clearly confirmed.

Reaction amplitudes for Ded1ΔC were significantly higher than with wtDed1 under

identical conditions (Fig.3.3E,F). This result corroborates the notion that Ded1 establishes an ATP-dependent steady state between unwinding and annealing reactions.

3.1.3 ATP modulates the balance between RNA unwinding and strand annealing.

To understand the mechanism by which Ded1 balanced the catalysis of the two

opposing processes, it was of interest to quantitatively assess the effect of the ATP

concentration on the ratio between unwinding and annealing rates. To this end, I

determined unwinding and annealing rate constants at different ATP concentrations

(Fig.3.4).

The unwinding rate constant increased with the ATP concentration, while the

annealing rate constant decreased (Fig.3.4A-C). This differential effect of the ATP

concentration on unwinding and annealing rate constants essentially enables Ded1 to

50 modulate its predominant activity in an ATP-dependent fashion. At low ATP concentrations, annealing is primarily catalyzed, at high ATP, unwinding is mainly promoted (Fig.3.4D). These results suggest that the ATP concentration is not only critical for establishing the steady state between unwinding and annealing but also for determining the ratio between the rate constants for both processes.

3.1.4 The balance between unwinding and annealing activities of Ded1 depends on

the RNA substrate.

To test whether the nature of the RNA substrate affected activities of Ded1, a

forked RNA complex containing unpaired nucleotides 5’ and 3’ to the duplex region of

the substrate used above (F[16], Fig.3.4E) was generated . Although the forked RNA

complex differed only in unstructured regions from the RNA used above (S[16]), time

courses at 2 mM ATP showed a significantly lower reaction amplitude (Fig.3.4E). (Note

that in all reactions, (i) Ded1 was present at saturating concentrations with respect to the

substrates, (ii) the concentrations of all substrates were held constant at 0.5 nM, and (iii)

the basal rates for uncatalyzed annealing were within the same order of magnitude for all

substrates.) Interestingly, the unwinding rate constant for F[16] was only slightly lower

than for S[16], whereas the annealing rate constant was significantly greater for F[16]

(Fig.3.4F). These findings suggest that the unstructured substrate regions significantly

impact the annealing activity of Ded1 while affecting the unwinding activity only

slightly.

Unwinding and annealing rate constants for F[16] over a range of ATP

concentrations (Fig.3.4G) were then determined. The unwinding rate constant increased

51 Figure3.4

S[16] F[16] S[19] 25 nt 16 bp 25 nt 16 bp 25 nt 19 bp A E I 25 nt 1 1 1 0.8 0.8 0.8 0.6 0.6 0.6 0.4 0.4 0.4 0.2 0.2 0.2 Fraction Unwound Fraction Fraction Unwound 0 0 Fraction Unwound 0 0 5 10 15 20 25 30 0 5 10 15 20 25 30 0 5 10 15 20 25 30 Time [min] Time [min] Time [min]

B F K

-1 -1 -1 -1 -1 -1 kunw = 3.3 min kann = 0.4 min kunw = 2.8 min kann = 3.4 min kunw = 0.5 min kann = 1.8 min (1.1·109 M-1min-1) (9.8·109 M-1min-1) (5.2·109 M-1min-1)

C G L ] ] ] -1 -1 5 -1 5 5 k 4 kunw 4 ann 4 3 3 3 kann 2 2 2 k 1 ann 1 kunw 1 kunw 0 0

Rate Constant Rate [min 0 Rate Constant [min Rate 0 0.5 1 1.5 2 [min Constant Rate 00.511.52 00.511.52 ATP [mM] ATP [mM] ATP [mM]

D H M

10 10 10 unw unw unw / k / k / k 1 1 1 ann ann k ann k k 0.1 0.1 0.1 0 0.5 1 1.5 2 0 0.5 1 1.5 2 0 0.5 1 1.5 2 ATP [mM] ATP [mM] ATP [mM]

Figure 3.4 Dependence of unwinding and annealing rate constants on ATP-concentration and the nature of the RNA substrate. (A) Representative unwinding timecourse for S[16] with 2 mM ATP. Reaction amplitude Aunw = 0.90 ± 0.05. (B) Rate constants for both unwinding and annealing reactions with S[16] at 2 mM ATP. The timecourse in panel (A) was fitted to eq. VI (Materials and Methods), yielding the indicated rate constants. (C) Dependence of unwinding (●) and annealing rate constants (○) on ATP concentration. Rate constants were determined as described in panel (B) from unwinding timecourses at each ATP concentration. Data points are the average value from multiple independent measurements, error bars

52 indicate one standard deviation. The curve for unwinding rate constants vs. [ATP] was fitted to a binding [max] -1 unw isotherm (eq. VII, Materials and Methods), yielding kunw = 4.4 ± 0.2 min and Km = 0.43 ± 0.04 mM. The line through the annealing rate constants vs. [ATP] represents a linear trend. (D) Ratio of unwinding and annealing rate constants vs. [ATP]. The ratio was calculated from the values given in panel (C). Note the logarithmic scale of the y-axis. The broken line emphasizes the ratio of one, values above the line indicate predominant annealing activity, values below the line predominant unwinding activity. The solid line represents a hyperbolical trend. (E) Representative unwinding timecourse for F[16] with 2 mM ATP (A = 0.45 ± 0.05). (F) Rate constants for both unwinding and annealing reactions with F[16] at 2 mM ATP. (G) Dependence of unwinding (●) and annealing rate constants (○) on ATP concentration for F[16]. Rate constants were determined as in panel (C). The curve for unwinding rate constants vs. [ATP] was fitted to a [max] -1 unw sigmoidal binding isotherm (eq. VIII), yielding kunw = 3.1 ± 0.3 min and Km = 0.36 ± 0.05 mM and a Hill coefficient of n = 2.2 ± 0.6. The broken line illustrates a hyperbolical binding isotherm (eq. VII). The line through the annealing rate constants vs. [ATP] represents a linear trend. (H) Ratio of unwinding and annealing rate constants vs. [ATP] for F[16]. The solid line represents a hyperbolical trend. (I) Representative unwinding timecourse for S[19] with 2 mM ATP (A = 0.24 ± 0.02). (K) Rate constants for both unwinding and annealing reactions with F[16] at 2 mM ATP. (L) Dependence of unwinding (●) and annealing rate constants (○) on ATP concentration for F[16]. Rate constants were determined as in panel (C). Unwinding rate constants vs. [ATP] were fitted to a hyperbolical binding isotherm (eq. VII), yielding [max] -1 unw kunw = 0.69 ± 0.04 min and Km = 0.55 ± 0.08 mM. (M) Ratio of unwinding and annealing rate constants vs. [ATP] for S[19]. The solid line indicates a hyperbolical trend.

53 with the ATP concentration to a similar extent as for S[16] (Fig.3.4G). The annealing rate

constant for F[16] decreased with the ATP concentration and the ratio between annealing and unwinding rate constants also decreased with the ATP concentration (Fig.3.4G,H).

To test how the length of the duplex region influenced the activities of Ded1, a

substrate where the duplex region of the complex was extended from 16 bp to 19 bp with

the single overhang (S[19], Fig.3.4I) was generated. Unwinding time courses at 2 mM

ATP showed a smaller amplitude than identical reactions with F[16] and S[16] (Fig.3.4I).

The unwinding rate constant was considerably lower than for the two other substrates,

whereas the annealing rate constant was greater than for S[16], yet smaller than for F[16]

(Fig.3.4K). The unwinding rate constant increased, while the annealing rate constant

decreased with increasing ATP concentrations (Fig.3.4L). The change in the ratio of

annealing and unwinding rate constants with S[19] was similar to the changes observed

with S[16] (Fig.3.4M). Notably, Ded1 catalyzed predominantly annealing with S[19] over

the entire range of ATP concentrations. However, increasing ATP concentrations

attenuated the annealing activity of Ded1 (Fig.3.4M).

Collectively, data with three different substrates demonstrate that the balance by

which Ded1 catalyzes disruption and formation of RNA duplexes strongly depends on the

nature of the substrate. In addition, the degree to which ATP modulates the ratio between

unwinding and annealing is influenced by the nature of the substrate. Our results reveal

the mechanistic cause for these observations: the unwinding rate constant decreases with

an extension of the RNA secondary structure whereas the annealing rate constant

increases with the overall length of the RNA strands, regardless of whether these are

present in structured or unstructured regions.

54 3.1.4 ADP also modulates the balance between RNA unwinding and strand

annealing.

While the data above clearly showed ATP-dependent modulation of the balance

between unwinding and annealing activities of Ded1, it remained unclear whether the

other prevalent cellular nucleotide phosphate, ADP, also affected the activities of Ded1.

ADP itself did not promote substrate unwinding, as shown above (Fig.3.1B). To ascertain

the effect of ADP on both unwinding and annealing rate constants, unwinding reactions

with the three substrates tested above in the presence of 2 mM ATP and increasing

amounts of ADP were conducted (Fig.3.5). The ATP concentration of 2 mM was chosen

in order to obtain significant reaction amplitudes for all substrates (Fig.3.4).

Amplitudes of unwinding reactions with S[16] decreased with increasing ADP

concentrations (Fig.3.5A). Both unwinding and annealing rate constants declined,

however, the relative decline of both rate constants differed (Fig.3.5B). Consequently, the

ratio between unwinding and annealing rate constants changed with increasing ADP

concentrations, i.e., annealing was favored over unwinding at higher ADP (Fig.3.5C).

These data indicate that the balance between unwinding and annealing can be modulated

by ADP.

The non-hydrolysable ATP analog AMPPNP did not alter the balance between

the two activities, even though increasing concentrations of AMPPNP inhibited both

unwinding and annealing activities (Fig.3.6A,B). Moreover, increasing concentrations of

free Mg2+ did not change the ratio between unwinding and annealing rate constants (Fig.

3.7A, B). Therefore, the balance between unwinding and annealing was altered

specifically by ADP and not by general inhibition of Ded1 by AMPPNP or Mg2+.

55 3.1.5 The degree of the ADP-dependent modulation between unwinding and

annealing depends on the RNA substrate.

The observed ADP-dependent alteration of the Ded1 activity with S[16] raised the

question whether the degree of this ADP-dependent modulation would also differ for

distinct substrates. To test this idea, unwinding timecourses with increasing ADP

concentrations with the substrates F[16] and S[19] were measured (Fig.3.5D,G). With F[16],

reaction amplitudes decreased only slightly with increasing ADP (Fig.3.5D). Both

unwinding and annealing rate constants declined (Fig.3.5E). Most importantly, the ratio

of both rates did not change as dramatically as seen with S[16] (Fig.3.5F). For S[19], reaction amplitudes for unwinding reactions declined much more than for F[16] with

increasing ADP (Fig.3.5G), indicating that unwinding of S[19] requires more ATP than the

substrates F[16] and S[16] with a shorter duplex region. Both unwinding and annealing rate constants decreased with increasing ADP and the ratio between unwinding and annealing rate constants changed with ADP to an extent similar to that seen with S[16] (Fig.3.5H,I).

AMPPNP did not alter the ratio between unwinding and annealing rate constants for

either F[16] and S[19] substrate to the extent of the effects observed with ADP (Fig.3.6).

These observations further establish that ADP, but not general inhibition of Ded1

by AMPPNP, can significantly alter the balance between unwinding and annealing rate

constants. As seen with ATP, the degree of this modulation strongly depends on the RNA

substrate.

56 Figure 3.5

S[16] F[16] S[19]

25 nt 16 bp 25 nt 16 bp 25 nt 19 bp A D G 25 nt 1 1 1

0.8 0.8 0.8

0.6 0.6 0.6

0.4 0.4 0.4

0.2 0.2 0.2 0

Fraction Single Strand 0 0 Fraction Single Strand Fraction Single Strand 0 5 10 15 20 25 30 0 5 10 15 20 25 30 0 5 10 15 20 25 30 Time [min] Time [min] Time [min] B E H

kunw k kann 1 1 ann 1

kunw kann 0.1 0.1 0.1 kunw

0.01 0.01 0.01 Rate Constant [min-1] Constant Rate [min-1] Constant Rate Rate Constant [min-1] Constant Rate 0 0.5 1 1.5 2 00.511.52 00.511.52 ADP [mM] ADP [mM] ADP [mM] C F I 2 4 15

1.5 3 10 unw unw unw k k k / / 1 / 2 ann ann ann 5 k k 0.5 k 1

0 0 0 0 0.5 1 1.5 2 0 0.5 1 1.5 2 0 0.5 1 1.5 2 ADP [mM] ADP [mM] ADP [mM]

Figure 3.5 ADP-dependent modulations of unwinding and annealing activities of Ded1. (A) Representative unwinding timecourses for S[16] in the presence of 2 mM ATP without ADP (●, reaction amplitude A = 0.90 ± 0.05), 1 mM ADP (○, A = 0.57 ± 0.02), and 2 mM ADP (♦, A = 0.39 ± 0.03). (B) Dependence of unwinding (●) and annealing rate constants (○) for S[16] on ADP concentration. Rate constants were determined as described in Fig.3.4B. (C) Ratio of unwinding and annealing rate constants vs. [ADP] for S[19]. Values were calculated as in Fig.3.4D. The line represents a linear trend. (D) Representative unwinding timecourses for F[16] in the presence of 2 mM ATP without ADP (●, A = 0.45 ± 0.05), 1 mM ADP (○, A = 0.40 ± 0.01), and 2 mM ADP (♦, A = 0.34 ± 0.01). (E) Dependence of unwinding (●) and annealing rate constants (○) for S[16] on ADP concentration. Rate constants were determined as described in Fig.3.4B. (F) Ratio of unwinding and annealing rate constants vs. [ADP] for F[16]. Values were calculated

57 as in Fig.3.4D. The solid line indicates a sigmoidal trend. (G) Representative unwinding timecourses for S[19] in the presence of 2 mM ATP without ADP (●, A = 0.24 ± 0.02), 1 mM ADP (○, A = 0.10 ±0.01), and 2 mM ADP (♦, A = 0.07 ± 0.01). (H) Dependence of unwinding (●) and annealing rate constants (○) for S[19] on ATP concentration. Rate constants were determined as described in Fig.3.4B. (I) Ratio of unwinding and annealing rate constants vs. [ADP] for F[16]. Values were calculated as in Fig.3.4D. The line represents a sigmoidal trend.

58 Figure 3.6

S[16] F[16] S[19]

25 nt 16 bp 25 nt 16 bp 25 nt 19 bp A C E 25 nt 1 1 1

0.8 0.8 0.8

0.6 0.6 0.6

0.4 0.4 0.4

0.2 0.2 0.2 0

0 0 Fraction Single Strand Fraction Single Strand Fraction Single Strand 0 5 10 15 20 25 30 0 5 10 15 20 25 30 0 5 10 15 20 25 30 Time [min] Time [min] Time [min] B D F 4 4 4

3 3 3 unw unw unw k k k / / 2 / 2 2 ann ann ann k k 1 k 1 1

0 0 0 0 0.5 1 1.5 2 00.511.52 00.511.52 AMPPNP [mM] AMPPNP [mM] AMPPNP [mM]

Figure 3.6 Effects of AMPPNP on unwinding and annealing activities of Ded1. (A) Representative unwinding timecourses for S[16] in the presence of 2 mM ATP without AMPPNP (●, A = 0.90 ± 0.05), 1 mM AMPPNP (○, A = 0.84 ± 0.06), and 2 mM AMPPNP (♦, A = 0.91 ± 0.08). (B) Ratio of unwinding and annealing rate constants vs. [AMPPNP] for S[16]. Values were calculated as in Fig.3.4D. The line represents a linear trend. (C) Representative unwinding timecourses for F[16] in the presence of 2 mM ATP without AMPPNP (●, A = 0.45 ± 0.05), 1 mM AMPPNP (○, A = 0.55 ± 0.10), and 2 mM AMPPNP (♦, A = 0.47 ± 0.06). (D) Ratio of unwinding and annealing rate constants vs. [AMPPNP] for F[16]. Values were calculated as in Fig.3.4D. The line represents a linear trend. (E) Representative unwinding timecourses for F[16] in the presence of 2 mM ATP without AMPPNP (●, A = 0.24 ±0.02), 1 mM AMPPNP (○, A = 0.29 ± 0.03), and 2 mM AMPPNP (♦, A = 0.27 ± 0.01). (F) Ratio of unwinding and annealing rate constants vs. [AMPPNP] for F[16]. Values were calculated as in Fig.3.4D. The line represents a linear trend.

59 Figure 3.7

A

1 0.8 0.6 0.4 0.2 0 Fraction Single Strand 0 5 10 15 20 25 30 Time[min] B

1

0.1

0.01

[min-1] Constant Rate 00.511.52 MgCl2 [mM] C 2

1 1.5 k

/ -1 k 1

Ratio 0.5

0 00.511.52

MgCl2 [mM]

Figure 3.7. MgCl2-dependent modulation of unwinding and annealing activities of Ded1. (A) [16] Representative unwinding timecourses for S in the presence of 2 mM ATP without MgCl2 (●, reaction

amplitude A = 0.90 ± 0.05), 1 mM MgCl2 (○, A = 0.83 ± 0.04), and 2 mM MgCl2 (♦, A = 0.76 ± 0.03). (B) [16] Dependence of unwinding (●) and annealing rate constants (○) for S on MgCl2 concentration. Rate constants were determined as described in Fig.4B. (C) Ratio of unwinding and annealing rate constants vs. [16] [MgCl2] for S . Values were calculated as in Fig.4D. The line represents a linear trend.

60 3.2 Discussion

It is shown here that Ded1 catalyzes two opposing processes in vitro, the

disruption and the formation of RNA duplexes. The balance between these two activities

is modulated by ATP and ADP concentrations. Moreover, the nature of the RNA

substrates significantly affects overall unwinding and annealing rates as well as the

degree by which ATP and ADP can modulate the balance between duplex formation and

unwinding. These findings expand the functional repertoire of DEAD-box proteins and

reveal the capacity of Ded1 to regulate RNA remodeling in response to ATP and ADP

concentrations and to features of the RNA substrate in an unexpectedly complex and

versatile fashion.

The ability of Ded1 to couple the distinct remodeling of different RNA substrates

to ATP and/or ADP concentrations is based on the capacity of the enzyme to establish an

ATP-dependent steady state between unwinding and annealing. Ded1 institutes this

steady state by catalyzing both unwinding and annealing in the presence of ATP. The rate

constants for both unwinding and annealing are sensitive to ATP and ADP concentrations. ATP “actively” promotes unwinding while suppressing annealing. ADP, on the other hand, inhibits both unwinding and annealing, yet to a different extent. The promotion of the unwinding activity by increased ATP concentrations most likely reflects the affinity of Ded1 for ATP during the unwinding process. ADP presumably interferes with ATP binding and thereby inhibits unwinding. A higher apparent affinity of the enzyme for ADP, compared to ATP, has been observed for other DEAD-box proteins as well (145, 147). Unexpected, however, are the effects of ATP and ADP on the annealing

activity of Ded1. Although more research is necessary to understand the mechanism of

61 ADP and ATP mediated inhibition of strand annealing, our data indicate that both ATP

and ADP impair the ability of Ded1 to facilitate strand-annealing (note that Ded1 was

present at saturating concentrations with respect to the RNA, even at the highest ADP

concentration used). The ADP bound state of Ded1 appears to be most unfavorable for

annealing, which, combined with the high affinity of Ded1 for ADP, is the reason why

ADP, but not AMPPNP can modulate the balance between unwinding and annealing rates.

Equally unexpected were the profound effects of structured and, notably,

unstructured regions in the RNA substrate on the balance between unwinding and

annealing rates. My data suggest that the remarkable sensitivity of Ded1 to the nature of

the RNA substrate is because unwinding and annealing are differentially affected by the

extent of structured regions and by the overall length of the RNA strands. The unwinding

rate constant decreases with the extent of the duplex region of the substrate, but is largely

insensitive to the overall length of the RNA. In contrast, the annealing rate constant

increases with the overall length of the RNA, regardless of whether the RNA is structured

or not. Thus, the impact of unstructured substrate regions on RNA remodeling by Ded1 is

conferred by the strand annealing activity of the enzyme.

The ability to catalyze both RNA unwinding and strand annealing is not a

universal feature of DEAD-box or DExH/D proteins, as demonstrated here (Fig.3.2) and

previously (164). Nevertheless, there are DEAD-box proteins that promote strand annealing in vitro, in addition to RNA unwinding (2, 4, 9, 11). Notably, these proteins

(human p68 and p72, yeast Mss116, and cyanobacterial CrhR) share with Ded1 a C- terminus that contains clusters of arginines and lysines. (Similarities among p68, p72,

62 Mss116, CrhR and Ded1 outside of this RK-rich segment do not exceed the level of

similarity to other DEAD-box proteins.) It is thus attractive to hypothesize that DEAD-

box proteins with C-terminal RK-clusters form a distinct subgroup of enzymes with the ability to catalyze both ATP-dependent RNA unwinding and strand annealing in vitro.

Consistent with this notion, the C-terminus of Ded1 is important, albeit not absolutely required, for the annealing activity of the enzyme (Fig.3.2). While a role for

RK-rich region in promoting duplex formation is a novel finding for DEAD-box proteins, other proteins with similar RK-rich segments, such as hnRNP A1 or hnRNP U, are known to facilitate strand annealing (200, 201). The exact role of RK-rich segments for

RNA strand annealing is presently unknown, but it may be possible that these RK-rich regions have similar functions in diverse proteins. The positively charged amino acids may neutralize charges of the sugar phosphate backbones of the RNA strands (202). Such charge neutralization is known to facilitate strand annealing (203).

Despite the prominence of the C-terminal RK-rich region, the strand annealing

activity of Ded1 does not exclusively reside in this protein segment (Fig.3.2, 3.3). Most

likely, catalysis of duplex formation by Ded1 also involves the helicase core domains

with the ATP binding site, which enables the enzyme to influence the annealing activity

through ADP and ATP ratios. An influence of ATP and ADP on the annealing activity

has also been observed for the cyanobacterial DEAD-box protein CrhR and the human

DEAD-box helicase Ddx42p(2, 4), suggesting that involvement of the helicase core

domains in the annealing activity is not unique to Ded1 (164). However, CrhR promotes

duplex formation only in the presence of ATP and Ddx42 displays higher annealing

activity with ADP than with ATP or without a nucleotide cofactor, whereas the annealing

63 activity of Ded1 decreases with increasing ATP or ADP concentration, indicating that

distinct DEAD-box proteins elicit different effects on their annealing activities from

ATP/ADP binding.

Interestingly, not only DEAD-box RNA helicases but also at least two DNA

helicases, RecQ5β and BLM, exhibit strand annealing activity in vitro (165, 166). The

annealing activity of RecQ5β also involves the C-terminus of the enzyme. As for Ded1,

the efficiency of duplex formation by RecQ5β and BLM decreases upon ATP binding.

The ability of RecQ5β and BLM to promote both duplex formation and unwinding shows that catalysis of these two opposing processes by the same enzyme is not restricted to “RNA helicases”. In fact, catalysis of duplex unwinding and strand

annealing by the same enzyme is not even unique to helicases. A diverse group of RNA

and/or DNA binding proteins, referred to as “RNA or DNA chaperones” also catalyze

both duplex unwinding and strand annealing (204). However, these “traditional”

RNA/DNA chaperones do not utilize NTPs or other co-factors and are therefore only able

to balance unwinding and annealing through protein concentration, i.e. through coverage

of their respective substrates(204). In contrast to these traditional RNA/DNA chaperones,

Ded1 uses the nucleotide co-factors to modulate the balance between duplex unwinding

and strand annealing. The ability of Ded1 to “tune” between duplex unwinding and strand

annealing in response to nucleotide co-factors, exceeds the functional capacity of

traditional RNA/DNA chaperones.

While the modulated remodeling of RNA structures through more than duplex

unwinding widens the horizon for possible biological roles of Ded1 (and perhaps other

related DEAD-box proteins), the physiological significance of combined unwinding and

64 annealing activities has not been explored for Ded1 or any other DEAD-box protein.

However, there are physiological examples of coordinated interactions between proteins

that facilitate duplex formation and DExH/D proteins that themselves do not promote

strand annealing. For instance, the DEAD-box protein eIF4A interacts with eIF4B, which

catalyzes strand annealing in vitro, and the DExH protein Brr2 functions in conjunction

with Prp24, which promotes annealing of RNAs (205-207). Although these examples do

not directly pertain to the physiological significance of concurrent unwinding and

annealing activities in a single DEAD-box protein, the eIF4A-eIF4B and Brr2-Prp24 interactions illustrate that proximity of helicase and strand annealing activities may be advantageous at certain points in RNA metabolism, such as during the ATP-driven exchange of mutually exclusive RNA-RNA interactions in mRNA splicing or ribosome biogenesis (28). A protein harboring both ATP-dependent unwinding as well as strand annealing activities would be well suited to catalyze such RNA strand exchanges without the need for other protein factors. Notably, Ded1 as well as p68 have been found in spliceosomal and ribosomal precursor particles, although physiological substrates for both proteins are presently unknown (185, 208).

Since DEAD-box proteins have also been implicated in displacing proteins from

RNA (23, 59, 61, 62, 209), it may be possible that enzymes such as Ded1 work as

protein-RNA exchangers. Such protein-RNA exchanges occur during spliceosome

function (28), although it is unknown whether Ded1 or p68 participate in these steps. It is

also important to note, that the intermolecular strand annealing activities shown for Ded1, p68 and other DEAD-box proteins, may in fact be relevant for intramolecular duplex

65 formation, i.e., RNA folding, as recently shown for the DEAD-box protein RH-II/Gu

(210).

Besides the interesting mechanistic possibilities afforded by DEAD-box proteins

that promote both strand annealing and disruption of RNA-RNA or RNA-protein

interactions, the regulation of the two opposing activities is of potential biological

importance. Because the balance between unwinding and annealing activities is sensitive

to ATP and ADP concentrations, it is conceivable that Ded1 utilizes ATP and ADP

modulation in one or more of its myriad of physiological roles (174). In fact, the maximal

modulation by ADP under our reaction conditions occurs in the range of physiological

ATP/ADP ratios (ATP concentrations: 2.2 – 9.1 mM, ADP concentrations: 0.6 – 2.9 mM,

ATP/ADP ratios: 2.47 – 9.25) (211, 212). It is thus possible that Ded1 adjusts its activity

under metabolic stress situations that are accompanied by changes in ADP or ATP concentrations. Interestingly, it has been shown that Ded1 genetically interacts with PAS kinase (188), a key enzyme involved in coordinated regulation of glucose flux, which is strongly influenced by changes in ADP and ATP levels, although numerous alternative explanations may equally well account for this observation (188).

In addition to the observed ADP- and ATP-dependent modulation of the Ded1

activities, posttranslational modifications such as phosphorylation or methylation may

change the dominant activity of the protein. I note that the alteration of RNA binding of

p68 through phosphorylation has been reported, although it is unknown whether the

modification also affects the balance between duplex unwinding and formation for this

enzyme (213, 214). Moreover, proteins containing RG-clusters are targets for

methylation (215). It will be interesting to investigate in vitro and in vivo whether

66 posttranslational modifications of DEAD-box proteins that catalyze both RNA unwinding

and annealing affect the balance between both activities.

Finally, the remarkably distinct response of Ded1 to different RNA substrates

may be physiologically significant. Ded1 has been implicated in numerous cellular processes such as mRNA splicing, RNA export and translation initiation, and it has been shown that the enzyme is required for translational regulation of specific genes (174,

216). An ability to differentially remodel distinct RNA substrates would certainly constitute a rational advantage for an enzyme with multiple biological functions.

67 Chapter 4: DEAD-box protein assisted RNA structure conversion towards and against thermodynamic equilibrium values

In the previous Chapter, I have showed that Ded1 facilitated the formation of

RNA duplexes, in addition to its ability to unwind duplexes. These observations suggested that Ded1 might also assist RNA structure conversions, i.e. interconversion of

RNA from one defined structure to another. RNA structure conversions occur frequently in biological processes such as pre-mRNA splicing, ribosome biogenesis, RNA export, and translation (28, 217-219). Examples for such RNA structure conversions (also called conformational switches) from the pre-mRNA splicing machinery include the exchange of U1 for U6, the disruption of the U4/U6 stem II and subsequent formation of the U6 3′ stem/loop, and the disruption of the U2 5′ stem/loop and subsequent formation of the

U2/U6 helix II (these and further examples are reviewed in ref (28)).

Even though RNA structure conversions share certain characteristics, many

questions remain about principles that govern RNA structure conversions. For example,

RNA structure conversions are frequently, if not always, assisted by proteins (28, 98,

204, 219, 220). However, the role of proteins in these complex structural changes of

RNA is not well understood. Furthermore, interconverting RNA structures are often

mutually exclusive, i.e. one RNA structure has to be disrupted to form a different one

(28). It is unclear how these structural arrangements are coordinated by proteins in an

accurate and well-timed fashion. In addition, more stable RNA structures are frequently

converted into less stable ones (28). Although this is a potentially important distinction to processes that govern the folding of functional RNAs, which generally occurs from less stable to more stable conformations (221), it remains uncertain how RNA structures can

68 be converted against thermodynamic equilibrium values and how proteins assist in such a reaction.

While the notion that Ded1 may assist RNA structure conversions is attractive, it is unknown how Ded1 may coordinate its unwinding and annealing activities to facilitate complex structural changes in RNA. Here, a simple model system is devised to recapitulate essential aspects of RNA structure conversions such as the mutually exclusive nature of the structures formed. This system is used to investigate how Ded1 facilitates complex structural changes in RNA in vitro.

69 4.1 Results

4.1.1 Substrate design

To understand Ded1-facilitated RNA structure conversions, a tripartite RNA

system was designed (strands x, y, z, Fig. 4.1). Strand y either formed a stable structure

with strand z (complex A), or with strand x (complex B) (Fig. 4.1A). That is, formation of

complex A required disruption of complex B, and vice versa. Complex B contained a binding site for the RNA binding protein U1A (62, 222), to test whether RNA structure

conversion could be coupled to subsequent protein deposition. For the substrates’

sequences, see table 4.1.

Complex B had a lower thermodynamic stability than complex A. At RNA

concentrations of 0.5 nM, the melting temperature for B was lower than that of A by 10°C

(Fig.4.1B). The stability difference was maintained with Ded1 bound; Ded1 bound to

complex A with a higher affinity (Kd = 17 ± 2 nM) than to complex B (Kd = 208 ± 17

nM). Both A and B were unwound by Ded1 in a strictly ATP-dependent fashion, and the

enzyme also facilitated the formation of both structures out of single strands (Fig. 4.2).

70 Figure 4.1

A A B

DED1 10 19 10

x * * y z

B 1 0.8

0.6 B A 0.4 Fraction Fraction 0.2 Strand Single 0 20 40 60 80 100

Temperature [°C]

Figure 4.1 Substrate design and characterization. (A) RNA substrate design. Three single strands (x, y, z) form two mutually exclusive structures (A, B). Numbers indicate the length of the basepaired region or the unpaired single stranded region. The asterisk indicates the position of the radiolabel. For sequences, see Supplementary Figure 1. (B). Thermal melting curves of structures A and B. The curves represent a A smoothed trend, yielding a melting temperature for complex A of Tm = 75.0 ± 0.5 ºC and a melting B temperature for complex B of Tm = 65.0 ± 0.5 ºC.

71 Table 4.1 Substrates and their sequences

Substrate Sequence

A (yz) 5’-CCCAGACAGCAUUGUACCCAGAGUCUGUACGG-3’ ||||||||||||||||||| CGUAACAUGGGUCUCAGAC-5’ 3’-UACAGUAACUACGACAAUCAUGCA

G U U U A A C A B (yx) 5’-CCCAGACAGC CCAG GUCUGUACGG-3’ |||||||||| |||| |||||||||| 3’-GGGUCUGUCG GGUC CAGACAUGCC-5’ C C A A U C U G

Tether 5’-CGCATACAGTCCACTGACGCAT –Biotin |||||||||||||||||||||| 3’- Connected to strand y Æ TTTTGCGTATGTCAGGTGACTGCGTA –5’

72 4.1.2 RNA structure conversion by Ded1

To measure the extent to which Ded1 could convert complex A into B, and vice

versa, it was necessary to determine the distribution of the two structures at the

thermodynamically controlled equilibrium under the respective reaction conditions. To

this end, pre-formed A (2 nM) was combined with strand x (8 nM). The mixture was

heated to 95°C and slowly cooled to reaction temperature, which allowed the two

structures to form in a distribution dictated by thermodynamic stabilities and respective

concentrations (Fig.4.3A). The resulting distribution of the RNA complexes was

analyzed on non-denaturing PAGE and yielded a distribution of 20 % B and 80 % A

(Fig.4.3A, lane ΔT). An identical distribution was detected when strand z was added to

pre-formed B (identical concentrations), indicating that 20 % B and 80 % A indeed

reflected equilibrium values (data not shown). Thus, as expected, the thermodynamic

equilibrium between the two RNA structures favored formation of the more stable A over

B by a factor of four at the reaction conditions and the concentrations used. Whether the

two RNA structures could interconvert without protein co-factor (Fig.4.3B) was then

tested. No notable conversion of A into B was detected within 6 h (Fig.4.3B), and no

conversion of B into A was seen when pre-formed B was incubated with strand z at concentrations and conditions identical to those for the reverse reaction (Fig.4.3C).

Addition of Ded1 without ATP resulted in a clearly measurable conversion of

complex A into B (Fig.4.3D). After transformation of 20 % A into B, the reaction reached a steady level that was virtually identical to the distribution between complexes A and B dictated by the thermodynamic equilibrium (i.e., 20 % B, 80 % A, Fig.4.3E). When complex B was pre-formed and conversion into A was measured, the same distribution

73 Figure 4.2

Annealing Unwinding A B *

* 0 30 0 30 Time [min] Time [min] C D

* * 0 30 0 30 Time [min] Time [min]

Figure 4.2 Ded1 can unwind complex A and B and anneal complex A and B from respective RNA strands. Reactions were performed at 24 ºC with 800 nM Ded1 and 0.5 nM RNA. Mobility of duplex and single stranded RNA is indicated by the cartoons on the left. Asterisks represent the radiolabel. (A) Representative PAGE of an unwinding time course of complex A. Zero represents the reaction before ATP addition. Aliquotes were removed between 0.5 min and 30 min. (B) Representative PAGE of a strand annealing time course with Ded1. No ATP was present during the reaction. Zero represents the reaction before Ded1 addition. Aliquots were removed from 0.5 min to 30 min. (C) Representative PAGE of an unwinding time course of complex B. (D) Representative PAGE of a strand annealing time course with Ded1.

74 Figure 4.3

A B D F H J L RNA only DED1 / no ATP DED1 / 0.1 mM ATP ATP depletion DED1 / 0.5 mM ATP A *

B * y ΔT 0 360 0 360 0 60 CS10 360 0 60 * Time [min] Time [min] Time [min] Time [min] Time [min] C E G I K 1 0.8 0.6 0.4 0.2 Fraction B Fraction 0 0 100 200 300 0 100 200 300 02040600 100 200 300 0204060 Time[min] Time[min] Time[min] Time[min] Time[min]

Figure 4.3 RNA structure conversions. (A) Size standards (left panel) and thermodynamically dictated equilibrium between structures A and B (right panel, ΔT). Cartoons indicate species A, B, and strand y, the asterisk represents the radiolabel. The green block arrow emphasizes that reactions were started with pre- formed complex A. The procedure to measure the thermal equilibrium is described in the Materials and Methods Section. (B) Representative timecourse of a structure conversion (A Æ B) with only RNA present. Aliquots were removed between 0 and 360 min. (C). Plots of timecourses for structure conversion A Æ B (●) and B Æ A (○). The fraction of complex B at any given time is plotted. The dotted line indicates the distribution between structures A (80 %) and B (20%) at the thermodynamically dictated equilibrium. (D) Representative timecourse of structure conversion A Æ B with Ded1 (800 nM). Aliquots were removed between 0 and 360 min. (E). Plots of timecourses for structure conversion A Æ B (●) and B Æ A (○). Lines represent the best fit to the integrated form of a homogenous first order rate law (ref). Observed first order

rate constants (kobs) and amplitudes (A) were computed from multiple independent experiments and were: AÆB -1 AÆB BÆA -1 BÆA kobs = 0.018 ± 0.003 min , A = 0.20 ± 0.01; kobs = 0.022 ± 0.003 min , A = 0.20 ± 0.01. (F) Representative timecourse of structure conversion A Æ B with Ded1 (800 nM) and 0.1 mM ATP. Aliquots were removed between 0 and 60 min. (G). Plots of timecourses for structure conversion A Æ B (●) and B Æ A (○). Lines represent the best fit to the integrated form of a homogenous first order rate law. Observed

AÆB rate constants and amplitudes were computed from multiple independent experiments and were: kobs = -1 AÆB BÆA -1 BÆA 0.48 ± 0.10 min , A = 0.61 ± 0.02; kobs = 0.45 ± 0.07 min , A = 0.60 ± 0.01. (H) Representative timecourse of approach to the thermodynamically dictated equilibrium after ATP depletion by hexokinase. Lane C represents the reaction (A Æ B) before addition of Ded1and ATP. Lane S indicates the reaction 10 min after the addition of Ded1 (800 nM) and 0.1 mM ATP; 10 indicates addition of 0.15 Units hexokinase and 1 mM glucose after these 10 min. Subsequently, aliquots were removed up to 360 min. (I) Plot of the timecourse after hexokinase addition. The line represents the best fit to the integrated hexo form of a homogenous first order rate law, yielding an apparent rate constant of kobs = 0.017 ± 0.006 min-1 and a final amplitude of A hexo = 0.24 ± 0.05. The two dotted lines represent the distribution of

75 structures A and B at the ATP dependent steady state (0.61) and at the thermodynamic equilibrium (0.20). (J) Representative timecourse of structure conversion A Æ B with Ded1 (800 nM) and 0.5 mM ATP. Aliquots were removed between 0 and 60 min. The yellow triangle indicates the mobility of the single strand y. (K). Plots of timecourses for structure conversion A Æ B (●) and B Æ A (○). Lines represent the best fit to the integrated form of a homogenous first order rate law. Observed rate constants and amplitudes

AÆB -1 AÆB were computed from multiple independent experiments and were: kobs = 2.2 ± 0.7 min , A = 0.61 ± BÆA -1 BÆA 0.02; kobs = 2.0 ± 0.6 min , A = 0.63 ± 0.02.

76 between A and B was reached (Fig.4.3E). These observations preclude the possibility that the levels of B and A were caused by formation of a dead-end complex. Rather, the data indicate that Ded1, in the absence of ATP, has the capacity to accelerate the rate constant by which the thermodynamic equilibrium between the two RNA structures was reached.

Next, the conversion of A into B by Ded1 in the presence of ATP was measured

(Fig.4.3F). The reaction proceeded significantly faster than the structure conversion without ATP, and, most importantly, an amplitude was reached at 60 % B (Fig.4.3F,G).

This level of B is clearly above the 20 % B seen in both the reaction by Ded1 without

ATP and in the establishment of the thermodynamic equilibrium between the two structures. Thus, Ded1 with ATP appeared to be able to convert a more stable RNA structure into a less stable one. Conversion of B into A with Ded1 and ATP also yielded

60 % B and 40% A, a distribution identical to that seen in the inverse reaction (Fig.4.3G).

This observation, again, precluded formation of dead-end complexes as a possible reason for the measured distribution. ATP alone did not facilitate any measurable structure conversion (data not shown). The non-hydrolyzable ATP analog AMPPNP also could not substitute for ATP (Fig 4.4), suggesting that hydrolysis and not mere ATP binding by

Ded1 were critical for converting the more stable RNA structure into the less stable one.

To further probe the role of ATP hydrolysis, the effect of ATP depletion during the reaction was tested. To this end, conversion of A to B with Ded1 and ATP was allowed to proceed for 10 min, after which the reaction amplitude of 60 % B was reached

(cf. Fig.4.3G). Then hexokinase (223, 224) was added, which hydrolyzed the ATP in the reaction mix within approximately 4 s (data not shown). I observed a clear decrease of the fraction B, accompanied by an increase in the fraction of A (Fig.4.3H). This transition

77 plateaued at 20 % B and 80 % A (Fig.4.3I), the distribution dictated by the thermodynamic equilibrium value (cf. Fig.4.3A). This result indicates that continuous

ATP hydrolysis by Ded1 is required to maintain a B/A distribution shifted towards the less stable structure.

In the presence of a higher ATP concentration (0.5 mM, Fig.4.3J), the distribution of B and A in the steady state was not significantly different from that at the lower ATP concentration (0.1 mM, Fig.4.3J,K). However, the reaction proceeded significantly faster at the higher ATP concentration (Fig.4.3K). These data show that greater energy expenditure during the reaction affected primarily the reaction kinetics and less the distribution between the RNA structures.

At the higher ATP concentration, I also noted a slight, yet significant accumulation of single stranded RNA (Fig.4.3J, triangle). A further rise in the ATP concentration to 2 mM resulted in an even larger fraction of this single stranded species

(Fig.4.3L, triangle). This observation suggested that the RNA structure conversion with

Ded1 and ATP involved an intermediate state of disassembled RNA.

78 Figure 4.4

DED1 + AMPPNP A

B

y ΔT 0 360 Time [min]

Figure 4.4 Representative timecourse of structure conversion A Æ B with 800 nM Ded1 and 0.5 mM AMPPNP, depicted as in Figure 4.3A. Aliquots were removed between 0 and 360 min.

79 4.1.3 Basic kinetic mechanism for ATP-dependent RNA structure conversion

The above results provided the basis to formulate a basic mechanism by which

Ded1 facilitates the RNA structure conversion in an ATP-dependent fashion. The single stranded RNA species emerging at higher ATP concentrations suggests that Ded1 uses

ATP-hydrolysis to disassemble the initial structures. Subsequently, Ded1 uses its annealing activity to facilitate formation of the final RNA structures (Fig.4.5). Fitting the time-courses for the RNA structure conversions with this simple two-step mechanism yielded rate constants that closely matched the respective rate constants that were separately determined for unwinding and annealing reactions of the respective complexes

(Fig.4.5). This basic mechanism quantitatively accounted (i) for the relative insensitivity of ATP concentrations from 0.1 mM to 0.5 mM of the distribution between A and B, (ii) for the increase of the rate constant at higher ATP concentrations, and (iii) for the larger fraction of single stranded RNA at higher ATP concentrations (2 mM).

Most importantly, the mechanism suggests that the kinetics of unwinding and

annealing, rather than thermodynamic stabilities of the RNA complexes determined the

distribution of the RNA structures. Ded1 establishes a kinetically controlled, ATP-

hydrolysis dependent, steady state between the two RNA structures, which allows a

distribution of B and A that is essentially independent of thermodynamic equilibrium

values. Removing hydrolyzable ATP results in a “return” of the system to the

thermodynamic equilibrium.

80 Figure 4.5 A A B B 0.25 min-1 / 0.1 mM ATP 1 1.1 min-1 / 0.5 mM ATP 2.4·109 M-1min-1 0.8 0.6 0.4 9 -1 -1 -1 5.6·10 M min 0.47 min / 0.1 mM ATP 0.2 2.1 min-1 / 0.5 mM ATP B Fraction 0 0204060 Time[min] Disassembled Strands

Figure 4.5. Basic kinetic scheme for structure conversion with Ded1 and ATP. (A) Rate constants were determined by numerically fitting timecourses in Fig.4.3 using the Kinsim/Fitsim software package (225).

Rate constants for unwinding of the complexes A and B at the given ATP concentrations were determined in separate unwinding reactions under identical conditions, as previously described (7). These rate constants were fixed during the fitting procedure and the annealing rate constants were computed. (B)

Simulated timecourses (lines) using the rate constants given in (a) for 0.5 mM ATP (gray) and 0.1 mM

ATP (black) for both structure conversions A Æ B (●), and B Æ A (○). The points included represent actual

data.

81 4.1.4 Formation of a tripartite RNA intermediate is critical for structure conversion without ATP

While a basic mechanism for the structure conversion with ATP had been illuminated, it remained unclear how Ded1 facilitated the reaction without ATP. The inability of Ded1 to unwind either RNA complex without ATP in the timeframe of our experiments (data not shown) precluded a mechanism involving intermediates of disassembled RNAs. If the strands were not disassembled, then, I hypothesized, there had to be intermediates that contained all three RNA strands in one complex. To probe the existence of such tripartite RNA species, a fluorescence technique was employed

(Fig.4.6A), in which strand x was labeled with Cy5 and strand z with Cy3. The two fluorophors undergo fluorescence resonance energy transfer (FRET) when they are within a distance of less than approximately 8 nm (226). Given the nucleotide separation of about 5 bp, representing a distance of ~ 1.2 nm in solution, a tripartite RNA species would be expected to provide a FRET signal 1.0 (Fig.4.6A).

When Cy3-labeled complex A was incubated with Cy5-labeled strand x, no FRET was observed (Fig.4.6B). I concluded that no significant accumulation of tripartite intermediates occurred with just RNA strands present. However, addition of Ded1 without ATP resulted in a small, yet significant FRET signal (Fig.4.6B). The FRET signal was strictly dependent on the presence of strand y, omitting this strand under otherwise identical conditions produced no FRET (Fig.4.6.C,D). The observed FRET therefore suggested a genuine tripartite species, not a Ded1-mediated aggregation of the strands z and x.

82 Figure 4.6

ABA FRET 1.0 Cy5 Cy3 RNA DED1 DED1 0.8 only ATP 0.6 Tripartite 0.4 Intermediate 0.2 Intensity [au] Intensity x 0 y z 550 600 650 700 550 600 650 700 550 600 650 700 Wavelength [nm]

CD 1.0 DED1 RNA DED1 0.8 only ATP 0.6 0.4 0.2 Intensity [au] Intensity 0 550 600 650 700 550 600 650 700 550 600 650 700 Wavelength [nm]

Figure 4.6 . Ded1- assisted stabilization of tripartite intermediate. (A) Scheme of formation of the tripartite intermediate. Position of the fluorescent labels. Cy3 is symbolized by a green circle (on complex A, strand z), Cy5 by a red circle on strand x. (B) Ensemble fluorescence spectra. Pre-formed complex A (Cy3) is incubated with strand x (Cy5) in the absence of Ded1 and ATP (left panel), with Ded1 (middle panel), and with Ded1 and 0.1 mM ATP (right panel). The triangle indicates the emission resulting from energy transfer of the tripartite complex. The FRET in the presence of Ded1 without ATP (middle panel) is approximately 0.04, as estimated by fitting of the spectra with three Gaussian curves and analysis of the areas corresponding to donor and acceptor emission after normalization to the spectra in the absence of Ded1. The FRET value with ATP (right panel) is ~ 0.02. (C) Scheme of formation of strand x and z aggregate in the absence of strand y, as depicted in panel A. (D) Ensemble fluorescence spectra. Strand z (Cy3) is incubated with strand x (Cy5) in the absence of Ded1 and ATP (left panel), with Ded1 (middle panel), and with Ded1 and 0.1 mM ATP (right panel).

83 Addition of 0.5 mM ATP decreased the FRET signal (Fig.4.6B), and the FRET signal disappeared at higher ATP concentrations (2 mM, data not shown). These results are consistent with the above finding that Ded1 disassembled the RNA strands in the presence of ATP.

FRET signals similar to those described above were also observed when Cy5 labeled complex B was incubated with Cy3 labeled strand z and Ded1 in the absence of

ATP (data not shown). Thus, the Ded1-facilitated formation of tripartite species was not restricted to strand x associating with complex A. Rather, a tripartite species could be formed as soon as an RNA region was available that allowed the pairing of a few bases

(Fig.4.6A). Collectively, these results demonstrate that without ATP Ded1 aids the formation of a tripartite intermediate.

However, the low FRET signal raised the question whether the tripartite intermediate was stable or only transiently formed. A low FRET signal could be caused by a highly populated (stable) state with low FRET efficiency, or alternatively, by a rarely populated, (unstable) state with high FRET efficiency (227-229). To distinguish between these two possibilities, fluorescence measurements on the single molecule level using the labeling scheme described above and a wide-field, TIR-based single molecule fluorescence detection system were conducted. (ref., Materials and Methods). Cy3- labeled complex A was immobilized and free, Cy5-labeled strand z was present in solution (Fig.4.7A). As in the ensemble measurement above, Cy3-labeled complex A was expected to show no FRET signal, whereas the tripartite should produce a higher FRET value.

84 Figure 4.7

A A FRET Cy5 Cy3 Tripartite Intermediate

x y z

B CY3Cy3

CY5Cy5 Intensity [au] Intensity

300 350 400 450 500 550 600 1

FRET 0 30 35 40 45 50 55 60 Time [s]

C 70 70 60 60 50 50 40 40 30 30 20 20 10 10 Number of Events of Number 0 of EventsNumber 0 0.0 0.2 0.4 0.6 0.8 1.0 0 0.5 1 1.5 2 FRET Time [s] Figure 4.7 . Measurement of Ded1- assisted stabilization of tripartite intermediate by single molecule FRET. (A) Scheme of formation of tripartite intermediate, as depicted in panel A, Figure 4.6. The grey diamond indicates the site at which complex A is immobilized to the surface for the single molecule measurements (Table 4.1, tether). (B) Formation of tripartite intermediate observed at the single molecule level. Representative timetrace of an individual complex A in the presence of Ded1 and Cy5 labeled strand x (upper panel). The yellow triangles emphasize the short, anti-correlated spikes of high Cy5 and low Cy3 intensity that are indicative of the transient formation of the tripartite intermediate. The lower panel shows the above timetrace converted to FRET intensity according to: FRET = Intensity(Acceptor) / Intensity(Acceptor) + Intensity(Donor) with intensities corrected for channel crosstalk (227). (C) Statistical analysis of FRET intensity (left panel) and duration (right panel) of the high FRET spikes. Individual spikes were identified in timetraces (panel B). The FRET intensity given in the left panel represents the average FRET value of the individual spikes, regardless of their duration. The number of spikes (y axis) with a given average FRET value (x axis) is plotted in the histogram. Duration of individual FRET spikes was measured with a customized Matlab program (228). The dwell time of molecules in the high FRET state (FRET > 0.6) is plotted in the histogram. The line represents a fit of the data points to a single

85 exponential decay, yielding τ = 0.17 ± 0.02 s and thus a rate constant by which strand x dissociates from the (x) -1 tripartite intermediate of kdiss = 353 ± 41 min .

86 In the presence of Ded1 without ATP, time trajectories showed long durations

without FRET (FRET ~ 0) punctuated by short spikes of high FRET (FRET ~ 1,

Fig.4.7B). In the absence of Ded1, only the low FRET state was observed and no FRET

spikes were detected in any of the time trajectories (data not shown). These observations

immediately suggested that in the presence of Ded1, the tripartite intermediate was

present only for short times. Analysis of FRET values for all recorded spikes revealed

that the FRET in these spikes was universally high, indicating that the intermediate

produced a high FRET signal (Fig.4.6B). A histogram of durations of the high FRET

spikes fit to a single exponential decay with a lifetime of τ = 0.17 ± 0.02 s, demonstrating

that the spikes were universally short (Fig.4.6C). Taken together, the fluorescence data

indicate that without ATP, Ded1 transiently stabilizes a tripartite intermediate that is

otherwise too fragile to accumulate.

With 0.5 mM ATP, Ded1 caused a significant decrease in the number of

immobilized complexes A over the course of a few min (data not shown). These

observations are consistent with the ability of Ded1 to unwind complex A in an ATP-

dependent fashion. In no case were the unwinding events accompanied by an increase in

FRET, strongly supporting the conclusion that, in the presence of ATP, Ded1 completely disassembled the RNA strands without formation of a tripartite intermediate. The overall decrease of the FRET signal due to ATP-dependent duplex unwinding was thus in excellent agreement with the ensemble FRET measurements, which showed a lower average FRET in the presence of these sub-saturating ATP concentrations (Fig.4.6B).

Time trajectories for Cy3-labeled complex A before unwinding events, however,

were highly similar to those without ATP (data mot shown). I concluded that these time

87 trajectories showed Ded1 without ATP bound, given the sub-saturating concentrations of

ATP.

4.1.5 Basic kinetic mechanism for RNA structure conversion without ATP

Having established the existence of a tripartite RNA intermediate for the Ded1 assisted structure conversion without ATP, I could now formulate a basic mechanism (Fig.4.8),

based on both fluorescence data (cf. Fig.4.6 & 4.7) and measurements of the RNA

structure conversions by PAGE (cf. Fig.4.3). The tripartite intermediate is only

transiently formed. The actual exchange of the RNA structures occurs from this tripartite

state, perhaps by a mechanism that may resemble branch migration (230-232). The two

RNA strands “equilibrate” according to their thermodynamic stability of the respective

complex, thus accounting for the observed approach to the thermodynamic equilibrium

value between the RNA structures in the presence of Ded1 without ATP. In fact, I

recorded several time trajectories of single complexes showing prolonged fluorescence

fluctuations, which would be consistent with “branch migration” events (Fig.4.9). The

number of trajectories showing these prolonged fluctuations was small (approximately 10

in ~ 104 time-trajectories measured), which precluded statistically sound analysis of any

putative “branch migration” events. It is important to note, however, that the kinetic

mechanism implies a slow step between the tripartite intermediates (Fig.4.8). Therefore,

the small number of trajectories showing the actual structure conversion is expected.

According to the kinetic mechanism, the tripartite intermediate should dissociate

approximately 1000 times before one complete structure conversion occurs, which is

consistent with the above observations.

88 Figure 4.8

ABA B 1 0.8 8 -1 -1 -1 -1 5.9·10 M min 0.2 min ~ 350 min 0.6 0.4 -1 -1 8 -1 -1 0.2

353 min 15.2 min ~5·10 M min B Fraction 0 0 100 200 300 Time[min]

Tripartite Intermediates

Figure 4.8 Basic kinetic scheme for Ded1-assisted structure conversion without ATP. (A) Rate constants for dissociation of strands x and z from the respective tripartite intermediates were determined with single molecule experiments as described in Figure 4.6D. Data for dissociation of z are not shown. Corresponding association constants (strand x and complex A, and strand z and complex B) were determined from the equilibrium between complexes A and B and the corresponding tripartite intermediates measured in the ensemble FRET experiments shown in Figure 4.6B. Data for ensemble FRET experiments of complex B with strand z are not shown. Rate constants for the actual strand exchange reaction (grey) were determined by numerically fitting timecourses in Fig.4.3D using the Kinsim/Fitsim software package (225). Association and dissociation rate constants that were determined separately (black numbers) were fixed and the strand exchange rate constants were allowed to float. (B) Simulated timecourses (lines) using the rate constants given in (a) for both structure conversions A Æ B (●), and B Æ A (○). The points included represent actual data.

89 Figure 4.9

Tripartite A Intermediates B

A B

CY3

CY5 Intensity [au] Intensity Intensity [au] Intensity

1 1 FRET FRET

0 0 60 80 100 120 024 6 seconds seconds

Figure 4.9 Possible branch migration intermediates during Ded1-assisted structure conversion without ATP. (A) Formation of the tripartite intermediate observed at the single molecule level. The upper panel shows the representative timetrace of an individual complex A in the presence of Ded1 and Cy5 labeled strand x. The lower panel shows the above timetrace converted to FRET intensity. (B) Interconversion between tripartite intermediates.

90 Regardless of the detailed mode by which the RNA structures exchanged, the

fluorescence data clearly showed that the RNA structure conversion by Ded1 without

ATP proceeded via a tripartite intermediate. This ATP independent pathway thus

contrasts with the reaction path taken with ATP, where the reaction proceeded via

disassembled RNA strands (cf. Fig.4.5).

4.1.6 Coupling RNA structure conversion to protein deposition

Despite the fundamental differences between the two pathways for the Ded1-

assisted RNA structure conversion, the kinetic data indicated that both reaction paths were highly dynamic processes of continuously changing RNA arrangements. It was thus pertinent to ask whether the complex B was at all sufficiently defined during the structure conversion to allow subsequent binding of the protein U1A, a well studied RNA-binding protein (222)(Fig.4.10A).

Complex B contained a U1A binding site that required formation of helices and

defined single stranded loops that were contributed by both strands y and x (Table 4.1),

i.e., U1A binding was only possible if complex B was formed to a significant extent

-3 during the reaction. Once bound, U1A dissociates only slowly from the RNA (kd = 10

min-1, (59)). Ded1 cannot accelerate the dissociation of U1A from the RNA (59). In addition, complex B with U1A bound is thermodynamically more stable than complex A

[U1A] (Kd = 5 nM, (59)). For these reasons, U1A binding was expected to cause an

accumulation of B exceeding the value observed during the reaction without U1A. No

additional accumulation of complex B during the RNA structure conversion in the

presence of U1A would indicate no U1A binding, and thus suggest only partial formation

91 of complex B during the structure conversion, or alternatively, Ded1 blocking the U1A

binding site.

When the structure conversion A Æ B with Ded1 and ATP was measured in the presence of U1A, almost complete conversion to complex B was observed (Fig.4.10B,C).

U1A could clearly bind during the reaction. RNA structure conversion with U1A and

Ded1 without ATP also yielded accumulation of complex B both faster and above the value seen with Ded1 alone. Thus, U1A also bound to complex B during the reaction without ATP. Taken together, these results show that complex B is formed to a significant extent during the reaction. Despite continuously changing RNA arrangements during RNA structure conversions, the RNA structure was sufficiently defined to enable binding of another protein that requires a rather extensive binding site (222).

Unexpectedly, and in contrast to the absence of measurable structure conversion

without any protein (cf. Fig.4.3B), conversion of complex A to B was observed with U1A

alone (Fig.4.10F,G). This finding suggests that U1A may interact with complex A, which

does not contain a U1A binding site. The U1A-facilitated structure conversion might

occur by a mechanism similar to that seen with Ded1 without ATP (cf. Fig.4.8).

92 Figure 4.10

[B - U1A] A A B

B D F * DED1 / ATP / U1A DED1 / U1A U1A only A *

BB * y * ΔT 0 120 0 120 0 120 Time [min] Time [min] Time [min]

C E G 1 0.8 0.6

0.4

Fraction B 0.2 0 0 40 80 120 0408012004080120 Time [min] Time [min] Time [min] Figure 4.10 Coupling of RNA structure conversion to deposition of U1A. (A) Reaction scheme. Complex -3 A was converted into complex B in the presence of U1A, which binds stably to B (Kd = 5 nM, kdiss = 10 min-1, (59)). (B) Structure conversion with Ded1, 0.5 mM ATP and U1A (120 nM), as depicted in Fig 4.3A. Structure conversions with U1A were measured as the reactions without protein, i.e., reactions were stopped with SDS (Materials and Methods); therefore no bands specific to any bound protein to complex B appear on the gel. (C) Plots of timecourses for structure conversion A Æ B with Ded1 and 0.5 mM ATP in the presence of U1A (120 nM) (○). Timecourse without U1A is shown as a reference (●). Data points represent the average of at least three independent experiments. Lines show the best fit to a single

AÆB [U1A] -1 exponential and yield, with U1A: kobs = 0.44 ± 0.05 min . (For data without U1A, see Fig.4.3). (D) Structure conversion with Ded1, 0.5 and U1A (120 nM) in the absence of ATP. (E) Plots of timecourses for structure conversion A Æ B with Ded1 and U1A (120 nM) without ATP (○). Data points represent the average of at least three independent experiments. Lines show the best fit to a double single exponential

I AÆB [U1A] -1 II AÆB [U1A] -1 and yield, with U1A: k obs = 1.68 ± 0.63 min , k obs = 0.0028 ± 0.0001 min (For data without U1A, see Fig.4.3E). (F) Structure conversion with only U1A (120 nM). (G) Plot of timecourse for structure conversion A Æ B with only U1A (120 nM). Data points represent the average of at least three

AÆB [U1A] independent experiments. The line represents the best fit to a single exponential yields: kobs = 0.0019 ± 0.0001 min-1.

93 4.2 Discussion

In this Chapter, a tripartite model system was used to investigate an RNA

structure conversion that is assisted by the DEAD-box protein Ded1. Despite its

simplicity, the model system recapitulates essential aspects of much more complex

physiological structure conversions. Notably the mutually exclusive nature of the two

RNA structures, the conversions towards, and, for the first time shown in vitro, against

thermodynamic equilibrium values, as well as the coupling of an RNA structure

conversion to the binding of another protein. Therefore, basic mechanistic features

elucidated in this work may also underlie more complex protein-assisted RNA structure

conversions.

It is shown here the Ded1-assisted RNA structure conversion can proceed via two

distinct pathways, depending on whether or not ATP is present. The pathway without

ATP involves a multipartite intermediate and results in a distribution of the RNA

structures dictated by thermodynamic equilibrium values. The ATP-dependent pathway

proceeds via completely disassembled RNA structures and allows RNA structure

conversion against thermodynamic equilibrium values. Although both pathways involve

continuously changing RNA arrangements, the RNA structures are sufficiently defined

during the structure conversion to enable binding of a protein with an extensive binding

site.

The mechanistic differences between the two pathways are best illuminated by a

free energy landscape of the RNA structure conversion (Fig.4.11). Stable reaction states

(i.e., stable RNA structures and stable intermediates) have low energy (blue depressions

94 Figure 4.11

A A + ATP -ATP

Tripartite Intermediate I Disassembled Strands Tripartite B Intermediate II

[B - U1A]

B

RNA only A

B [B - U1A]

A Tripartite DED1 Intermediate I

Tripartite Intermediate II B [B - U1A]

ΔΔG [kcal·mol-1] DED1 + ATP A 16

12 Disassembled Strands 8

B [B - U1A] 4 0

Figure 4.11 Ded1-assisted RNA structure conversion. (A) Schematics of the ATP-dependent (left) and the ATP-independent (right) pathway of the structure conversion. Complexes and intermediates are labeled as in the preceding text and figures. (B) Energy landscape of the ATP-dependent and ATP-independent pathways. Blue depressions correspond to the ground state of the indicated complexes. The ground state energy of complex A was taken as the zero reference value. The ground state energy of complex B was calculated from the distribution of complexes A and B in the thermodynamic equilibrium (Fig.4.3A), the ground state energy of the B-U1A was determined by adding the free energy of U1A binding to complex B U1A-B ‡ (Kd = 5 nM, ref. (59)). Transition state energies (ΔG ) correspond to observed rate constants given in ‡ the kinetic schemes in Figure 3 and 5, and were calculated according to ΔG = RT ln (h·kobs / kB·T),B where h

95 is Planck constant, kB B Boltzmann constant (233). Coloring of the landscape corresponds to the respective transition state energy, the color legend is given on the right besides the lower panel. The landscapes were generated with Matlab. White arrows indicate the preferred path of the RNA structure conversion without ATP (middle panel) and in the presence of ATP (lower panel). Note that the energies given in a landscape apply only at the given concentrations of RNA and ATP. Alterations in RNA concentrations change ground state energies, alterations in ATP concentrations change transition state energies for the ATP-dependent structure conversion pathway.

96 the reaction did not occur with the non-hydrolysable ATP analog AMPPNP (data not

shown). Unwinding of substrates with a DNA loading strand in the absence of the ability

in the surface). The height of the energy barrier between two neighboring states describes

the likelihood and thus the rate constant for a transition between these two states. If the

energy barrier is high, the rate constant for the transition is low and vice versa.

In the absence of protein co-factors, the height of the energy barriers between

both RNA structures (A and B) and the state where the strands are completely disassembled preclude a structure conversion via this pathway within the time of observation (Fig 4.11A). Without protein co-factors, I also did not observe structure conversion via tripartite intermediates (cf. Fig.4.3) or accumulation of tripartite intermediates within the detection time (cf. Fig.4.6&4.7). Therefore, neither of the two pathways for RNA structure conversion can be sufficiently populated without protein co- factors, and no interconversion between the two RNA structures occurred during the reaction time (cf. Fig.4.3B).

If Ded1 is present together with ATP, the RNA strands are completely

disassembled. Subsequently, Ded1 uses its annealing activity to re-form the other duplex.

The ATP-driven duplex unwinding by Ded1 lowers the energy barrier between the initial

RNA structures and the completely disassembled state, which opens the pathway for the

structure conversion (Fig.4.11B, lower panel). This pathway constitutes a kinetically

controlled steady state between unwinding and annealing reactions: the ratio between the

RNA structures is solely determined by the rate constants for unwinding and annealing

reactions, not by the thermodynamic stability of the structures. Therefore, the ability to establish a kinetically controlled steady state between the two RNA structures enables

97 Ded1 to facilitate an RNA structure conversion against thermodynamic equilibrium

values. To our knowledge, this is the first time such a process has been directly demonstrated in vitro. Continuous ATP hydrolysis by Ded1 is necessary to maintain a ratio of RNA structures against their thermodynamic equilibrium values. In this respect, the function of Ded1 resembles ATP-driven ion pumps, which maintain ion gradients against thermodynamic equilibrium values (234). In our model system, Ded1 “pumps” a more stable RNA conformation into a less stable RNA conformation in an ATP- dependent fashion.

The Ded1-assisted RNA structure conversion without ATP does not involve

completely disassembled strands but tripartite intermediates, which were directly

observed (Fig.4.6&4.7). Our data suggest that that Ded1 stabilizes these species.

Although the intermediates still dissociated most of the time, for every 1000 dissociation

events now approximately one transition from one intermediate to the other occurred

(Fig.4.8). This small probability is apparently sufficient to enable the interconversion of

the two RNA structures within the reaction time.

It remains unclear how exactly the transition between tripartite intermediates and final RNA structures occurs. Our observations suggest a process resembling a branch migration. The few single molecule trajectories that presumably show such events

(Fig.4.9) imply that the branch between the two strands moves back and forth, which would explain equilibration of the RNA structures according to their respective thermodynamic stability. Our data do not indicate whether Ded1 (without ATP) accelerates the branch migration process, but I note that branch migrations can occur spontaneously, as has been observed in other systems (235, 236).

98 Irrespective of the actual mode of the branch migration, our results show that

RNA structure conversion can be facilitated through “simple” stabilization of inherently

unstable RNA complexes. Conceivably, stabilization of the tripartite intermediate

correlates with the capacity of Ded1 to facilitate strand annealing (7). In fact, acceleration of strand annealing by a protein may be based on the stabilization of inherently unstable

species that form during the rate limiting steps of duplex formation (237, 238). Implicitly,

an enzyme that scores in an annealing assay might in fact function to stabilize inherently unstable RNA structures, which may explain why many RNA-binding proteins facilitate strand annealing (219, 239).

In this regard it is worth noting that even the RNA-binding protein U1A induced

the RNA structure conversion in our model system (Fig.4.10). U1A also accelerates the

bimolecular rate constant for strand annealing by one order of magnitude in a non-

sequence specific fashion (Q.Y . and E.J., unpublished results). These observations

support the above assertion that “RNA-binding proteins” may also facilitate RNA

structure conversions and thus carry out “functions” beyond simple association with

designated RNA binding sites. It may be interesting to test whether other RNA binding

proteins are able to perform similar “functions” in other systems and, most importantly,

in the cell.

As the pathway for the Ded1-facilitated RNA structure conversion without ATP

only involves equilibrium binding steps, the final distribution of the RNA structures is dictated by the thermodynamic equilibrium value. In this respect, this structure conversion pathway resembles strand exchange reactions catalyzed by proteins with RNA chaperone activity such as Ncp7 that also produce a ratio of RNA structures dictated by

99 thermodynamic equilibrium values (204, 219, 239). Most proteins with RNA chaperone

activity have significant annealing activities, which, as discussed above, may correlate

with a capacity to stabilize inherently unstable RNA species (240). It is thus attractive to

speculate that proteins with RNA chaperone activity may also facilitate RNA structure

conversions via the stabilization of tripartite intermediates, although it remains to be

directly shown whether this pathway is in fact taken by these proteins.

The kinetics for the two RNA structure conversion pathways illuminate features

advantageous for proteins that function in such processes. Proteins that facilitate RNA structure conversions without ATP would be efficient when they stabilize tripartite intermediates to a high degree and/or if they accelerate the subsequent “branch migration”. Proteins that employ ATP hydrolysis would be more efficient if the strand separation is fast, although this rate would perhaps need to be coordinated with the subsequent strand annealing; otherwise single stranded species would accumulate, which may or may not be problematic. However, the ATP-dependent pathway may be significantly faster than the ATP-independent pathway. Thus, employing ATP-driven

RNA unwinding activities in addition to annealing activities for RNA structure conversions not only allows RNA structure conversions against thermodynamic equilibrium values but also allows those reactions to occur rapidly. Potentially, fast

structure conversions allow better timed conformational changes in RNPs.

100 Chapter 5 The DEAD-box protein Ded1 unwinds RNA duplexes by a mode distinct from translocating helicases

In the previous Chapter, I showed that Ded1 could coordinate its annealing and unwinding activities to facilitate complex structural changes in RNA. These results suggested a prominent role for the unwinding activity of Ded1 even in the context of complex RNA structure rearrangement. In this Chapter, I investigate the molecular mechanism by which Ded1 unwinds RNA duplexes.

It has been proposed, and in some cases shown, that helicases translocate in one

direction along one strand of DNA or RNA in an ATP-driven manner, thereby displacing

either complementary nucleic acids (helicase activity) or bound proteins (25, 156, 157,

162, 241-243). To unwind duplexes, helicases are thought to first bind to one of the two

nucleic acid strands, the loading strand, and then unidirectionally translocate on this

loading strand through the duplex, which causes strand separation (155). Since the initial

binding for most helicases requires a single stranded loading region (153, 154), many

helicases that have been mechanistically studied exclusively unwind substrates

containing a single stranded region in one strictly defined orientation, either 5’or 3’

relative to the duplex (153-155). Helicases translocating in the 5’ to 3’ direction are

thought to require the single strand 5’ to the duplex, those translocating 3’ to 5’ are

assumed to need the single strand 3’ to the duplex.

However, most, if not all, DEAD-box helicases studied so far (105, 148), unwind duplexes regardless of the orientation of single stranded regions (51, 52, 58). While these results raised questions about unidirectional helicase translocation on the loading strand as a universal mode for duplex unwinding by all helicases (130, 153), it has remained

101 unclear how strand separation can occur otherwise. To understand how DEAD-box helicases accommodate single stranded regions of either orientation during the unwinding of RNA duplexes, the unwinding mode of Ded1 is investigated.

102 5.1 Results

5.1.1 Ded1 unwinds RNA duplexes without strict polarity.

Ded1 unwound RNA duplexes with either 5’ or 3’ single stranded regions, i.e.,

without strict polarity (Fig.5.1A,B). In addition, Ded1 unwound blunt-end duplexes,

although this reaction proceeded with a significantly slower rate (Fig.5.1C). These

observations showed that the lack of unwinding polarity by Ded1 was not due to fast

unwinding of duplexes from the blunt side of the duplex, as seen with the RecBCD DNA helicase complex (244, 245). Supplementing single stranded RNA in trans did not enhance the unwinding rate of the blunt-end duplex, suggesting that proximity between single strand and duplex had to be physically maintained (Fig.5.2).

Unwinding reactions of the above substrates with increasing concentrations of

Ded1 showed that the two substrates with the single stranded regions could be saturated

with Ded1 at similar concentrations (Fig.5.1D,E). The sigmoidal shape of these

functional binding curves suggested that Ded1 operates as an oligomer. At enzyme

saturation, the substrate with the 3’ overhang was unwound 2-fold faster than that with

the 5’ overhang, presumably due to small differences in duplex stability conferred by the

dangling nucleotides of the different overhangs (7). However, other mechanistic reasons

might account for this observation.

In contrast to the substrates with the single stranded regions, the blunt-end

substrate could not be saturated with Ded1 at experimentally accessible concentrations

(Fig.5.1F). This observation explained the significantly reduced unwinding rate for this

substrate and suggested that the single stranded regions aided the binding of Ded1. This

notion was further supported by equilibrium binding measurements in the presence of the

103 Figure 5.1

3’ 5’ blunt A B C * * *

* * *

0 5 0 5 0 45 Time [min] Time [min] Time [min] D E F 4.0 4.0 4.0 ]

-1 3.0 3.0 3.0

2.0 2.0 2.0 . [min

1.0 1.0 1.0 unwind k 0 0 0 0 200 400 600 800 0 200 400 600 800 0 200 400 600 800 DED1 [nM] DED1 [nM] DED1 [nM]

Figure 5.1 Ded1 unwinds RNA duplexes irrespective of the orientation of single stranded regions. For sequences see Table 5.1. (A)Representative PAGE of an unwinding time course for a 16 bp duplex with a 25 nucleotide single stranded overhang 3’ to its duplex region (3’oh). Mobilities of duplex and single stranded RNAs are indicated by the cartoons on the left, asterisks indicate the radiolabel. The zero time point represents the reaction before ATP addition. Aliquots were removed between 15 s and 5 min. (B) Representative PAGE of an unwinding time course for a 16 bp duplex with a 25 nucleotide single stranded overhang 5’ to its duplex region. (C) Representative PAGE of an unwinding time course for a 16 bp blunt- end duplex (blunt). Aliquots were removed between 30 s and 45 min. (D) Dependence of the unwinding rate constants of the 3’oh substrate on Ded1 concentration. Data points are the average values from multiple independent measurements. The curve for unwinding rate constants vs. [Ded1] was fitted to a

[max] -1 unw sigmoidal binding isotherm, yielding kunw = 3.3 ± 0.2 min , Kd = 144 ± 11 nM, and a Hill coefficient of n = 5.9 ± 2.5. Unwinding rate constants are likely to reflect contributions from multiple reactions steps. (E) Dependence of unwinding rate constants of the 5’oh substrate on Ded1 concentration.

[max] - The resulting curve was fitted to a sigmoidal binding isotherm as above, yielding kunw = 1.2 ± 0.2 min 1 unw , Kd = 129 ± 7 nM, n = 5.3 ± 1.2. (F) Dependence of unwinding rate constants of the blunt-end substrate on the Ded1 concentration. Data points are the average value from multiple independent measurements. The line connecting the data points represents a trend. The observed unwinding rate

-1 constant at 600 nM Ded1 was kunw = 0.1 min .

104 Table 5.1 Substrates and their sequences used in Figure 5.1. Cartoons correspond to those in Figure 5.1.

Substrate Sequence

5’-AGCACCGUAAAGACGC-3’ 3’ |||||||||||||||| 3’- AAAACAAAACAAAACAAAACAAAAUUCGUGGCAUUUCUGCG-5’

5’ 5’- AAAACAAAACAAAACAAAACAAAAUAGCACCGUAAAGACGC-3’ |||||||||||||||| 3’- UCGUGGCAUUUCUGCG-5’

5’- AGCACCGUAAAGACGC-3’ blunt |||||||||||||||| 3’- UCGUGGCAUUUCUGCG-5’

105 Figure 5.2

A

*

B 0.14

] -1 0.12

[min

0.10

unwind.

k 0.08 0 50 100 150 200

Molar Excess Single-Stranded RNA

Figure 5.2 Supplementing single stranded RNA in trans does not enhance the basal unwinding rate constant of the blunt-end substrates. (A) Reaction scheme. Single stranded RNA was added to the unwinding reaction with radiolabeled (asterisk) blunt-end duplex used in Figure 5.1. (B) Dependence of the unwinding rate constant of blunt-end RNA (0.5 nM) on the concentration of a 25 nucleotide single stranded RNA. Rate constants were determined from unwinding time courses at each concentration of the single stranded RNAs, as described in Figure 5.1. Ded1 was present at 600 nM. Data points are average values from at least two independent measurements, the line represents a trend.

106 non-hydrolyzable ATP analog AMPPNP (246). At a Ded1 concentration of 600 nM,

stable complexes were formed only with the substrates containing the single stranded regions, but not with the blunt-end duplex (Fig.5.3). Nuclease protection indicated that

Ded1 in fact bound to the single stranded regions (data not shown). Collectively, these data indicated that single stranded regions facilitated both functional and equilibrium binding of the enzyme.

5.1.2 Ded1 unwinds substrates with DNA loading strands

To elucidate how single stranded regions of either orientation promoted duplex

unwinding by Ded1, I employed substrates where one or both strands consisted of DNA

(Fig.5.4). Ded1 did not unwind DNA-DNA substrates (Fig.5.4A,B), although it was able

to bind these substrates in the presence of AMPPNP (Fig.5.4C). As previously seen by

others (246), Ded1 separated duplexes with an RNA loading and a DNA top strand

(Fig.5.4A,B). Other RNA helicases have also been shown to displace DNA from an RNA

loading strand (24, 51, 67, 72, 197). Ded1 unwound the RNA-DNA substrates with a

greater rate constant than their more stable RNA-RNA counterparts, consistent with the

previous observation that duplex stability affected the rate constant for strand separation

(7).

Unexpectedly and in contrast to other RNA helicases, Ded1 efficiently displaced

RNA from a DNA loading strand (Fig.5.4A,B). Unwinding rate constants for these

substrates were at least as high as for RNA-RNA substrates. This fast unwinding of

substrates with a DNA loading strand required a single stranded region (Fig.5.4D), and

and the reaction did not occur with the nonhydrolyzable ATP analog AMP-PNP (data not

107 Figure 5.3

DED1 -+ + DED1 -+ + DED1 -+ + AMPPNP --+ AMPPNP --+ AMPPNP --+ RNA RNA RNA DED1 DED1 DED1

RNA RNA RNA

Figure 5.3 Equilibrium binding of different substrates by Ded1. (A) Representative PAGE of equilibrium binding of 600 nM Ded1 to 0.5 nM 3’oh substrate without (middle lane) and with 1 mM AMPPNP (right lane). The arrows on the left indicate the position of the free substrate and Ded1-RNA complex. (B) Representative PAGE of equilibrium binding of Ded1 to the 5’oh substrate. Conditions were as in panel A. (C) Representative PAGE of equilibrium binding of Ded1 to the blunt-end substrate. Conditions were as in panel A.

108 Figure 5.4

A B 5′ 3′ R R R R D D D D D R R R D D D R 024 6 8 10 024 6 8 10 −1 −1 kunwind [min ] kunwind [min ] C D

R DNA R D DED1 D R D DNA D R D D 0 100 800 024 6 8 10 −1 DED1 (nM) kunwind [min ] E

3′ 5′ 5′ 3′ D R 3′ 5′ 024 6 8 10

−1 kunwind [min ]

Figure 5.4 Unwinding of RNA-DNA hybrid substrates by Ded1. For sequences see Table 5.2. (A) Unwinding rate constants (sliding points) of substrates with 3’overhangs. Gray lines indicate DNA, black lines RNA strands. Unwinding rate constants were determined as described for Fig.5.1D, Ded1 was present at saturating concentrations. Rate constants above 10 min-1 were not experimentally accessible with high accuracy and should be considered lower limits of the actual rate constants. (B) Unwinding rate constants of substrates with 5’overhangs. (C) Equilibrium binding of Ded1 (100 – 800 nM) to 0.5 nM the radiolabeled DNA substrate the 3’ overhang. Binding reactions were conducted as described in the Materials and Methods section in the presence of AMPPNP, except that no single stranded RNA was added prior to the loading of the samples on PAGE. Mobilities of free DNA and Ded1-DNA complex are indicated on the left. (D) Unwinding rate constants of blunt-end substrates in the presence of 600 nM Ded1. (E) Unwinding rate constant for a substrate with DNA bottom strand and a single stranded region of inverted polarity. Respective ends are indicated, the bottom strand is covalently connected through a 5’-5’ link. Arrows indicate 3’Æ 5’ polarity. Reactions were performed as in panel A.

109 Table 5.2 Sequences and characterization of substrates used in Figure 5.4

-1 Substrate Sequence Kd [nM] kunw [min ]

5’-AGCACCGUAAAGACGC-3’ |||||||||||||||| 144 ± 11 3.3 ± 0.2 3’- AAAACAAAACAAAACAAAACAAAAUUCGUGGCAUUUCUGCG-5’

ab 5’-AGCACCGTAAAGACGC-3’ -3 |||||||||||||||| N.A. ≤ 10 3’- AAAACAAAACAAAACAAAACAAAATTCGTGGCATTTCTGCG-5’

5’-AGCACCGTAAAGACGC-3’ c c |||||||||||||||| N.A.A. ≥ 10 3’- AAAACAAAACAAAACAAAACAAAAUUCGUGGCAUUUCUGCG-5’

5’-AGCACCGUAAAGACGC-3’ |||||||||||||||| 389 ± 12 3.4 ± 0.1 3’- AAAACAAAACAAAACAAAACAAAATTCGTGGCATTTCTGCG-5’

5’- AAAACAAAACAAAACAAAACAAAAUAGCACCGUAAAGACGC-3’ |||||||||||||||| 129 ± 7 1.2 ± 0.1 3’- UCGUGGCAUUUCUGCG-5’

5’- AAAACAAAACAAAACAAAACAAAATAGCACCGTAAAGACGC-3’ a b |||||||||||||||| N.A. ≤ 10-3 3’- TCGTGGCATTTCTGCG-5’

5’- AAAACAAAACAAAACAAAACAAAAUAGCACCGUAAAGACGC-3’ |||||||||||||||| 305 ± 16 3.9 ± 0.2 3’- TCGTGGCATTTCTGCG-5’

5’- AAAACAAAACAAAACAAAACAAAATAGCACCGTAAAGACGC-3’ d d |||||||||||||||| N.A.A. ≥ 10 3’- UCGUGGCAUUUCUGCG-5’

5’- AGCACCGUAAAGACGC-3’ e e |||||||||||||||| > 3,000 0.1 3’- UCGUGGCAUUUCUGCG-5’

5’- AGCACCGTAAAGACGC-3’ ab |||||||||||||||| N.A. ≤ 10-3 3’- TCGTGGCATTTCTGCG-5’

5’- AGCACCGUAAAGACGC-3’ e e |||||||||||||||| > 3,000 0.1 3’- TCGTGGCATTTCTGCG-5’

5’- AGCACCGTAAAGACGC-3’ e e |||||||||||||||| > 3,000 0.1 3’- UCGUGGCAUUUCUGCG-5’

Cartoons correspond to those in Figure 5.4, RNA is black and DNA is gray. Functional dissociation constants (Kd) and unwinding rate constants (kunw) were determined for each substrate as shown in Figure 5.

1D. Unwinding rate constants represent the values at enzyme saturation, unless indicated otherwise. (a) Kd

for the DNA-DNA substrates could not be determined because unwinding of this substrate by Ded1 was

undetectable. (b) No significant strand separation for the DNA-DNA substrates was detected after 60 min,

which sets the indicated upper limit for the unwinding rate constant. (c) Strand separation by Ded1

110 proceeded faster than experimentally accessible with high accuracy. After 5 s, more than 80 % of the final amplitude was reached. While this value sets the indicated lower limit for the unwinding rate constants,

lack of accurate information about the actual unwinding rate constant at enzyme saturation precludes exact

calculation of the corresponding functional dissociation constants Kd. However, experiments conducted at

lower temperature (data not shown) where the reactions proceed more slowly, in conjunction with

experiments where aliquots were taken faster than 5s (data not shown), allow for an estimate of the

constants at the given reaction temperature. These extrapolations yield: Kd ~ (130 ± 30) nM, kunw ~ (32 ± 8) min-1. I emphasize that values obtained from these experiments are less accurate than parameters determined directly from experiments at the actual reaction temperature and with aliquots taken every 5 s

or slower. (d) Strand separation by Ded1 proceeded faster than experimentally accessible with high

accuracy. After 5 s, more than 80% of the final amplitude was reached. The indicated value is thus a lower

limit for the unwinding rate constant. Extrapolating the corresponding data as described in (c) yields: Kd ~

-1 (190 ± 40) nM, kunw ~ (24 ± 7) min . (e) The blunt-end substrates could not be saturated with Ded1 (Figure

1f). The values for Kd thus represent lower limits. The values for kunw are unwinding rate constants at 600 nM Ded1.

111 shown). Unwinding of substrates with a DNA loading strand in the absence of the ability

to unwind DNA-DNA substrates is inconsistent with unidirectional helicase translocation

along the loading strand as a prerequisite for strand displacement.

If Ded1 were to translocate along a DNA loading strand during duplex unwinding,

DNA-DNA substrates should be separated as well. In addition, the RNA top strand was also released when the polarity of the single stranded region on the DNA was inverted

(Fig.5.4E). This result further supported the assertion that Ded1 does not unwind duplexes by a unidirectional translocation on the loading strand starting from the single stranded region.

5.1.3 Ded1 unwinds substrates without single strand–duplex junction.

If single stranded regions did not serve as the start sites for translocation, then

how did these substrate parts promote functional binding and duplex unwinding by

Ded1? There are two possibilities: either the junction of single strand and duplex was

critical for fast duplex unwinding, or that the single stranded region was required

proximal to the duplex to enable Ded1 to act directly on the helix. To distinguish between these two possibilities, a series of multi-piece substrates (MPS) that did not feature a single strand – duplex junction were designed. In the first MPS, a labeled DNA and a

labeled RNA formed a duplex adjacent to each other with a piece of unlabeled DNA

(Fig.5.5). The labeled DNA piece contained an unpaired DNA region, whereas the RNA

formed a blunt-end duplex with the unlabeled DNA. In the presence of Ded1 and ATP,

the RNA piece was efficiently released, but, as expected, not the DNA piece (Fig.5.5A,

left panel, lane 3). Notably, the RNA was unwound with a rate constant comparable to

112 Figure 5.5

3′

D R D R A **3′ **3′ 3′ B D 5′ D 5′ DED1 - + + + - + + + NPH-II - + + + - + + + ATP - - + + -- + + ATP --+ + --+ + 95 °C - -- + - --+ 95 °C ---+ ---+

* * * * * * * * * *

* R R * * * R R * * * * 1 2 3 4 1 2 3 4 1 2 3 4 1 2 3 4 Figure 5.5. Unwinding of multi-piece substrate I (MPS I) by Ded1 but not by NPH-II. For sequences see table 5.3 (A) MPS I. Gray lines, DNA; black lines, RNA; asterisks, radiolabels. Left panel: representative unwinding reaction of MPS I. Mobilities of the individual species are indicated by the cartoons on the left. Presence of Ded1 (200 nM, lane 2) and ATP (lane 3) in the reaction is indicated above the gel. Reactions were performed for 10 min. 70 % of the RNA substrate was unwound (lane 3). Right panel: representative unwinding reaction of the MPS I lacking the single stranded region. Reaction conditions were identical to those shown on the left panel, in the reaction with ATP (lane 3), 4 % of the RNA was unwound. (B) No release of the RNA piece from MPS I by NPH-II. Left panel: Representative unwinding reaction of MPS I. Reactions were performed for 5 min with 40 nM NPH-II (saturating with respect to duplex substrates with single stranded overhangs (25). Under these reaction conditions, NPH-II unwinds duplexes with 3’ overhangs with a rate constant of ≥ 3 min-1 (ref. (25) and data not shown). The slight DNA displacement is consistent with a previously shown slow DNA strand separation activity that is distinct from the pronounced RNA unwinding activity of NPH-II (197). Right panel: representative unwinding reaction of the MPS I lacking the single stranded region. Reaction conditions were as above.

113 Table 5.3 Sequences of multi-piece substrates (MPS). Cartoons correspond to those in

Figure 5.5 and 5.6. Gray lines, DNA; black lines, RNA

Substrate Sequence

AAAACAAAACAAAACAAAACAAAA-3’ 3’ T 5’-ACGAGGGAGACGAGGAAGCACCGUAAAGACGC-3’ 3’ * * |||||||||||||||||||||||||||||||| 5’ 3’-TGCTCCCTCTGCTCCTTCGTGGCATTTCTGCG-5’

3’ 3’ 5’-AGCACCGUAAAGACGCAGCAGGGAGACGAGGA-3’ ** |||||||||||||||||||||||||||||||| 5’ 3’-TGCTCCCTCTGCTCCTTCGTGGCATTTCTGCG-5’

3’ 5’-AGCACCGUAAAGACGCAGCAGGGAGACGAGGA-3’ ** |||||||||||||||||||||||||||||||| 3’ 5’ TGCTCCCTCTGCTCCTTCGTGGCATTTCTGCG-5’ T AAAACAAAACAAAACAAAACAAAACAAAACAAAACAAAAAC-3’

5’-biotin-UAAAACAAAACAAAACAAAACAAAACAAAA-3’ 3’

5’ 5’- AGCACCGUAAAGACGC-3’ * |||||||||||||||| 3’- UCGUGGCAUUUCUGCGUU-biotin-5’

114 that seen with substrates with a “regular” DNA overhang (data not shown). Removing the

single stranded region from DNA largely prevented the release of the RNA piece

(Fig.5.5A, right panel). These results clearly showed that a single stranded region on one

DNA piece promoted unwinding of a neighboring RNA piece. Therefore, a single strand

- duplex junction was unlikely to be necessary for fast unwinding by Ded1.

In addition, the data lend further support to the notion that Ded1 does not

translocate along the loading strand to unwind duplexes. To address this issue more

directly, whether an RNA helicase that unwinds duplexes by translocating along the

loading strand would then fail to release the RNA piece from the MPS was tested. To this end I examined if the RNA piece was removed by the viral DExH helicase NPH-II

(26, 27), which translocates 3’ to 5’ on the loading strand (25, 156). As expected, no significant release of the RNA was observed (Fig.5.5B, left panel, lane 3). This result directly suggested that Ded1 unwinds duplexes in a manner distinct from the translocating NPH-II.

To confirm and extend these observations, a second MPS where a labeled DNA

and a labeled RNA bound adjacent to each other on a piece of unlabeled DNA was

designed (Fig.5.6). The unlabeled DNA contained a 41 nucleotide long single stranded

region extending from the side opposite the RNA-DNA duplex (Fig.5.6). Ded1 readily

unwound the RNA, but not the DNA piece (Fig.5.6A, left panel). Lack of the single

stranded region on the DNA prevented the rapid release of the RNA piece (Fig.5.6A, right panel). These data showed that the single stranded region only had to be within a certain proximity, not in the immediate vicinity of the unwound duplex to facilitate strand separation by Ded1. In addition, the results confirmed that only the presence of a single

115 Figure 5.6

D R D R A **3′ **3′ 3′ B 3′ D 5′ D 5′

DED1 - + + + - + + + NPH-II - + + + ATP - - + + - - + + ATP - - + + 95 °C - - + - - + - - 95 °C - - - +

* * * * * * * * *

R R R * * * Figure 5.6 * * * 1 2 3 4 1 2 3 4 1 2 3 4 Unwinding of multi-piece substrate II by Ded1 but not by NPH-II. For sequences see table 5.3. (A) MPS II. Left panel: representative unwinding reaction of MPS II. Mobility of the individual species is indicated by the cartoons on the left. Reactions were performed for 5 min with 600 nM Ded1. With ATP (lane 3), 75 % of the RNA was released. Right panel: representative unwinding reaction of MPS II lacking the single stranded region. Reaction conditions were identical to those shown on the left panel. With ATP (lane 3), 8 % of the RNA was released. (B) No release of the RNA piece from MPS II by NPH-II. Representative unwinding reaction of MPS II. Reactions were performed as in Figure 5.5B. The corresponding unwinding reaction of MPS II lacking the single stranded region was identical to that shown in Figure 5.5B.

116 stranded RNA on the nucleic acid complex, but not a single strand – duplex junction on

the unwound duplex was necessary for fast duplex unwinding by Ded1. Finally, release

of the RNA piece without displacement of the DNA piece between the single strand and

the RNA piece lend additional support to the claim that Ded1 did not translocate from the

single stranded region to unwind duplexes. The translocating DExH helicase NPH-II

failed to release the RNA piece from this MPS (Fig.5.6B), further emphasizing

differences between the unwinding modes of Ded1 and NPH-II.

Next, it was examined whether proximity of single strand and duplex had to be

established by a nucleic acid linkage or whether single strand and duplex could also be

brought close by a protein link. To this end, a MPS where both duplex and single stranded RNA were modified with biotin was designed, and proximity between duplex and single strand was maintained through the biotin-binding protein streptavidin

(Fig.5.7A).A proximal single stranded RNA significantly stimulated unwinding of the duplex (Fig.5.7A, left panel). No enhancement of the basal unwinding rate constant of the duplex was seen without the single strand (Fig.5.7A, right panel), or without streptavidin

(Figure 5.8). These observations indicate that only proximity between the duplex and

single stranded regions is critical, not a specific type of linkage. The results further

showed that Ded1 unwinds the duplex by directly acting on the helical region. The single

stranded region apparently facilitates loading of Ded1 onto the duplex, and this loading

requires proximity between duplex and single stranded region. This unwinding mode

clearly differs from the translocation-based unwinding mechanism used by NPH-II.

Consequently, NPH-II was unable to unwind the duplex with or without the proximal

strand present (Fig.5.7B).

117 Figure 5.7

A 3′ 3′ B 5′ * 5′ * 3′

DED1 - + + + - + + + NPH-II - + + + - + + + ATP - - + + - - + + ATP - - + + - - + + 95 °C - - - + - - - + 95 °C - - - + - - - +

* * * * * * * * 1 2 3 4 1 2 3 4 1 2 3 4 1 2 3 4

Figure 5.7. Unwinding of the multi-piece substrate III by Ded1 but not by the DExH RNA helicase NPH-II. For sequences see table 5.3. Grey circles, biotin; the patterned shape, streptavidin. (A) Left panel: representative unwinding reaction of MPS III with Ded1. Mobilities of the individual species are indicated by the cartoons on the left. Reactions were performed for 5 min with 800 nM Ded1 (0.5 nM duplex, 2.5 nM streptavidin, 20 nM single stranded RNA). With ATP (lane 3), 69 % of the duplex was unwound. Right panel: representative unwinding reaction of MPS III without the single stranded RNA. With ATP (lane 3), 15 % of the duplex was unwound. (B) Left panel: representative unwinding reaction of MPS III with the DExH helicase NPH-II. Reactions were performed for 5 min with 40 nM NPH-II (saturating with respect to duplex substrates with single stranded overhangs (197)), 0.5 nM duplex, 2.5 nM streptavidin, 20 nM single stranded RNA. Right panel: representative unwinding reaction of MPS III without the single stranded RNA.

118 Figure 5.8

3’

5’* 5’*

DED1 - + + + - + + + ATP --+ + --+ + 95°C ---+ ---+

*

*

Figure 5.8 Unwinding of MPS III components without streptavidin. Reactions were conducted exactly as described in Figure 5.7, except that streptavidin was omitted from the reaction. The radiolabeled substrate in the left panel was unwound to 20 % after 5 min (third lane from left, 600 nM Ded1, and compare Figure 5.7), in the right panel the substrate was unwound to 14 %, under identical conditions. Thus, unwinding of the duplex was not enhanced by the single stranded RNA as observed in the presence of streptavidin.

119

5.2 Discussion

It is shown here that the DEAD-box RNA helicase Ded1 can unwind a

remarkably diverse set of substrates. As long as one piece of RNA is present in the

duplex region, Ded1 will unwind duplexes. This helicase activity is greatly stimulated by

single stranded RNA or DNA regions, which can be covalently attached to the duplex on either end and in either orientation (Figs.5.1,5.4). Moreover, the unwinding activity by

Ded1 is stimulated by single strands that are not covalently connected to the duplex

(Figs.5.5&5.6). Most notably, unwinding is even stimulated by a single strand that is

brought near the duplex by a streptavidin-biotin link (Fig.5.7).

These observations are inconsistent with the unwinding mechanism assumed to

underlie duplex unwinding by most helicases: binding of the enzyme at a single strand adjacent to the duplex and subsequent unidirectional translocation along this initially bound (loading) strand, away from the single stranded region (155). It is well established that such translocation-based helicase activity requires a single strand in one exclusive orientation with respect to the duplex. In fact, helicases are frequently classified based on these substrate needs as either 5’ to 3’, or 3’ to 5’ helicases. Unwinding of substrates with single stranded regions in either orientation (Fig.5.1), displacement of RNA but not DNA from DNA bottom strands (Fig.5.4A,B), and unwinding of a substrate where single stranded overhang and duplex region of the loading strand have opposite polarities

(Fig.5.4E), are all incompatible with a translocation-based unwinding mode for Ded1. In addition, helicases translocating from single stranded regions can not normally unwind substrates without covalent linkage between single strand and duplex, as directly shown

120 here for NPH-II (Figs.5.5B, 5.6B, 5.7B), and previously demonstrated for another

translocating RNA helicase, HCV-NS3 (157). However, I note that the DNA helicase complex RecBC, which translocates on one strand but does not start at an unpaired overhang, has been shown to “step” over covalent gaps in the loading strand (247). While

RecBC thus formally accommodates substrates without covalent linkage in the loading stand, unwinding only occurs when the loading strand pieces are in a particular collinear orientation, properly spaced, and bound to the same DNA strand (247). This coordinated, processive stepping of RecBC contrasts with Ded1, which clearly does not require specific orientations or arrangements between single strand and duplex for unwinding. I emphasize that I do not question that unidirectional helicase translocation underlies duplex unwinding by many, if not most DNA helicases, or that RNA helicases such as

HCV-NS3 and NPH-II unwind duplexes by translocation along the loading strand.

However, Ded1, and, as outlined below, possibly numerous other DEAD-box proteins, employ a distinct unwinding mode.

Our data indicate that Ded1 utilizes a single stranded region not as start site for unidirectional translocation but to facilitate the loading of the enzyme onto the duplex,

where strand separation is directly initiated. While strand separation by Ded1 can be

observed for substrates without single stranded regions (Fig.5.1C), functional and

equilibrium affinity of Ded1 for these substrates is low (Fig.5.1F, Fig.5.3C), indicating

that Ded1 can not form stable complexes with substrates lacking single stranded regions

(Fig.5.3C). However, Ded1 binds stably to substrates with single stranded regions

(Fig.5.3A,B, and Fig.5.4C). Bringing a single strand in the vicinity of a duplex enables

Ded1 to interact more frequently, or more productively with the duplex, which increases

121 unwinding efficiency, compared to substrates without single stranded regions

(Figs.5.1,5.4-5.7). As directly shown with the multi-piece substrate in Figure 5.7, single stranded regions, even if not covalently attached to the helix, aid loading of Ded1 onto the duplex.

How this “loading” occurs remains to be elucidated. It is possible that one functional unit of Ded1, which can be a monomer or an oligomer, is transferred from the single stranded region to the duplex, without leaving the nucleic acid. Alternatively, different functional units of Ded1 may bind single stranded and duplex regions.

Preliminary evidence points towards this scenario (cf. Chapter 2). It is also not yet clear whether a single strand allosterically activates duplex unwinding by Ded1, i.e., whether the single strand enhances the rate constant for strand separation, although preliminary data suggest that this is not the case (cf. Chapter 6). However, the single strand may induce conformational changes in Ded1 that enable tighter binding to the duplex. On the other hand, the single stranded region could simply increase the local concentration of

Ded1 around the duplex, which might lead to an increase in the number of productive duplex binding events, which might reflect the sigmoidal character of the unwinding rate constants over the Ded1 concentrations. While further experiments are required to illuminate these points, it appears clear that the facilitated loading of Ded1 is not a highly tuned process, since many different means of placing single strand proximal to the duplex region enhance Ded1’s unwinding activity.

As observed here for Ded1, several other DEAD-box proteins have also been shown to unwind RNA duplexes irrespective of the orientation of single stranded regions

(51, 52, 58). It is thus tempting to speculate that the mode of duplex unwinding employed

122 by Ded1 applies to other DEAD-box proteins as well. Since DEAD-box proteins are the

largest subgroup of RNA helicases (163), duplex unwinding without translocation on the loading strand may be prevalent in RNA metabolism. In conjunction with the limited or absent unwinding processivity of most DEAD-box proteins, many of which may only perform one unwinding step (148), our results raise the possibility that many DEAD-box proteins may not function as directional motors, but as “strand separation switches”.

Recent structural studies on the DEAD-box proteins, VASA and eIF4AIII, support this

view (75, 77, 131). Unwinding RNA structures by loading the helicase directly onto the

duplex and then “switching” the two strands apart, may be well suited for enzymes that

encounter mostly short duplexes within large RNP assemblies (28, 106). In addition, an

unwinding mode where the strand separating enzyme does not have to move extensively

on the RNA appears well tuned towards the strictly local conformational changes in

RNA/RNP assemblies that DEAD-box proteins are thought to catalyze in the cell (28,

106).

Even though duplex unwinding by Ded1 does not involve translocation of the

enzyme along the loading strand, it is important to note that Ded1 has been shown to

displace proteins from RNA without duplex unwinding (61). While it is unlikely,

although certainly not impossible, that this protein displacement involves actual

translocation of Ded1 on the RNA (61), our data now show that the ability of an enzyme

to perform ATP-driven conformational work on single stranded RNA does not predict

that duplex unwinding is based on movement along the loading strand as well. For Ded1,

protein displacement from single stranded RNA and duplex unwinding are two

mechanistically distinct processes.

123 Perhaps most unexpectedly, our results reveal the existence of at least two fundamentally different modes of duplex unwinding within the highly conserved family of DExH/D RNA helicases (106). Ded1, and presumably other DEAD-box proteins, unwind duplexes without translocating on the loading strand. In contrast, and as mentioned above, the viral DExH helicases NPH-II and HCV NS3 clearly function by unidirectional translocation along the loading strand (25, 156, 241, 248). Translocation- based unwinding, which resembles the mechanism thought to apply to many DNA helicases, may enable viral DExH RNA helicases such as NPH-II to unwind duplexes much longer than those separated by DEAD-box proteins (27, 106). The differences in the unwinding modes of DEAD-box vs. DExH proteins also raise the interesting prospect that the signature DEAD-box vs. DExH may predict the unwinding mode. Although this remains to be shown, the substrates used here provide a simple way to test whether a

DExH/D protein unwinds duplexes with or without translocation on the loading strand.

124 Chapter 6 Duplex unwinding by DEAD-box helicases from both terminal and internal helical regions

In the previous Chapter, I showed that Ded1 used a single stranded overhang not for starting translocation but for loading onto the helical region, where strand separation

was directly initiated. This novel unwinding mode raised two significant questions. First,

does the overhang have a role beyond loading, i.e., does the overhang influence the actual

strand separation? Second, is strand separation initiated at a particular terminus of the

helical region to initiate unwinding?

To answer these questions, I studied duplex unwinding by two DEAD-box

helicases Ded1 and Mss116. These two enzymes were chosen because they show robust

unwinding activity but are phylogenetically distant within the DEAD-box helicase family

(9).

125 6.1 Results

6.1.1 The overhang does not affect the unwinding rate constants of the strand

separation.

First, the effect of the overhang length on unwinding efficiency of Ded1 was

investigated. To this end, I devised a series of substrates containing sequentially extended

overhangs (5 nt – 35 nt) 3’ to a 16 bp duplex (Fig.6.1). Unwinding reactions were

performed for each substrate with increasing Ded1 concentrations. Unwinding rate

constants were recorded and plotted vs. Ded1 concentrations, yielding a functional

affinity for each substrate (Fig.6.1). A blunt-end substrate, as well as the substrate

containing a 5 nt overhang, could not be saturated at experimentally accessible Ded1

concentrations (Fig.6.1). The functional affinity of Ded1 for the substrate with a 15 nt

′ ′ overhang was Kd = 436 nM (Fig.6.1). The functional affinities (Kd ) of Ded1 for the

′ substrates with 25 nt and 35 nt overhangs were Kd ~ 200 nM (Fig.6.1). At saturating concentrations of Ded1, unwinding rate constants for the substrates with 15 nt, 25 nt and

35 nt overhang were virtually identical, with the maximal unwinding rate constants of

~3.3 min-1 (Fig.6.1). Collectively, these data suggest that the overhang affects the

loading efficiency but does not influence actual strand separation. These findings are consistent with our previous data that suggest a loading function for the overhang (cf.

Chapter 5). Indeed, it has been shown that a blunt-end substrate was unwound with a rate similar to that for an overhang substrate under saturating Mss116 concentrations, supporting the idea that the overhang did not affect the strand separation (9).

126 Figure 6.1

16 bp 4.0 ]

35 nt -1 3.0 25 nt 15 nt 2.0

5 nt 1.0 Constant [min Constant blunt Unwinding Rate 0.0 0 200 400 600 800 1000 DED1 [nM]

Figure 6.1 Dependence of unwinding rate constants on increasing Ded1 concentrations for 16 bp RNA substrates containing a 35 nt (♦), 25 nt (□), 15 nt (■), or 5 nt (○) single stranded RNA region at the 3’-end or without a single stranded region (●) as shown in the cartoons on the left. Unwinding rate constant at each Ded1 concentration was obtained from corresponding unwinding time course and represented the average values of at least two independent measurements. The lines through the data points represent a best

-1 ′ fit to a Hill equation, yielding k max(15) = 3.4 ± 0.1 min , K d(15) = 436 ± 11 nM, and a Hill coefficient of n

-1 -1 ′ (15) = 3.4 ± 0.3 min for substrate with a 15 nt overhang; k max (25) = 2.7 ± 0.9 min , K d(25) = 198 ± 27 nM,

′ and n (25) = 3.7 ± 0.2 for that with a 25 nt overhang; k max (35) =3.4 ± 0.1, K d(35) = 204 ± 22 nM, and n (35) =

2.7 ± 0.7 for that with a 35 nt overhang. The lines through the data points for substrates with a 5 nt or

without a ssRNA represent a trend.

127 6.1.2 Unwinding by Ded1 can be initiated from both termini of the helical region.

Next, it was asked whether Ded1 initiates strand separation at a specific terminus

of the helical region. To answer this question, I took advantage of the fact that Ded1

unwound RNA/DNA duplexes with a RNA top strand and a DNA loading strand but not

DNA/DNA duplexes (249). I reasoned that successively decreasing the RNA portion of

the top strand should point to the respective locations for unwinding initiation. First, I

prepared a series of chimeric top strands, where I substituted a portion (6 nt) of the RNA

strand with DNA either at 5’- or 3’-end of this strand. Next, I annealed these chimeric

strands to a 41 nt DNA bottom strand (Fig.6.2A), yielding RNA/DNA chimeric

substrates containing a 6 bp DNA/DNA region covalently linked to a 10 bp RNA/DNA

region (Fig.6.2A).

Ded1 unwound both chimeric substrates with the 5’- or 3’-end DNA substitution

(Fig.6.2A). Notably, chimeric substrates with the 3’-end DNA substitution were unwound

at least 6-fold faster than those with the 5’-end DNA substitution (Fig.6.2A). Removal of

the overhang prevented fast unwinding of both chimeric substrates (Fig.6.2B), indicating that the efficient unwinding of chimeric substrates much as substrates with a complete

RNA top strand required the presence of the overhang. These data suggest that Ded1 can initiate unwinding from both 5’- and 3’-RNA portion of the helical region. Upon opening the RNA/DNA part of the duplex, the remaining 6 bp DNA/DNA duplex most likely dissociates spontaneously, leading to the complete separation of the duplex.

Alternatively, Ded1 might be able to act on the DNA/DNA duplex after initiation of

Ded1 on the RNA/DNA duplex. In any event, RNA is required for initiation of unwinding and this initiation can occur on the either side of the duplex.

128 Figure 6.2

AB R5′ 3′ R5′ 3′ D D 5′ 3′ 5′ 3′ 5′ 3′ 3′ 5′ D R 3′ 5′ 3′ 5′ 0.1 1 10 -1 3′ 5′ k unwind [min ] 3 5 ′ ′ 0.1 1 10 -1 k unwind [min ]

Figure 6.2 Unwinding of RNA-DNA chimeric substrates by Ded1. Gray lines, DNA; black, RNA. 5’ and 3’ represent the 5’- and 3’-end of the top strand. Note that the DNA piece is covalently connected to the RNA piece on the chimeric strand. For sequences, see table 5.1. (A) Unwinding rate constants (sliding points) of 16 bp RNA-DNA chimeric substrates with a 25 nt DNA overhang at the 3’- or 5’-end. Unwinding rate constants were determined as in Fig.6.1. Ded1 was present at saturating concentrations. (B) Unwinding rate constants of blunt-end duplexes. Unwinding rate constants were measured with 600 nM Ded1 and the unwinding reactions for blunt-end duplexes could not be saturated at experimentally accessible Ded1 concentrations.

129 6.1.3 Unwinding by Ded1 does not require a free RNA terminus.

The finding that Ded1 could initiate unwinding on the either side of the duplex

raised the possibility that Ded1 might need a free RNA terminus for strand separation. To

test this idea, I designed a multiple-piece substrate (MPS) with a labeled 16 nt

RNA/DNA chimeric oligonucleotide and a labeled 30 nt DNA oligonucleotide binding

adjacent to each other on a unlabeled 71 nt DNA oligonucleotide. The RNA terminus at

the 5’-end of the labeled chimeric piece was blocked by the labeled DNA piece

(Fig.6.3A). Ded1 efficiently unwound the chimeric piece but, as expected, not the DNA

piece (Fig.6.3A). The chimeric piece was unwound slightly slower than regular chimeric

substrate (Fig.6.3B), which might be caused by increased stability of chimeric duplex in

the MPS through the coaxial helix stacking between the chimeric piece and the adjacent

DNA piece. Nonetheless, the data clearly show that blocking the RNA terminus does not

prevent unwinding, suggesting that Ded1 does not require a free RNA terminus for strand

separation.

6.1.4 Ded1 initiates unwinding within the helical region.

If Ded1 does not require a free RNA terminus, it should be able to initiate unwinding within the helical region. To test this hypothesis, I designed a top strand with

3 nt DNA at each end of 10 nt RNA and annealed this chimeric strand to the 41 nt DNA

bottom strand used before (Fig.6.2A, Fig 6.4A). Ded1 readily unwound this substrate

(Fig.6.4A). Unwinding reactions with increasing concentrations of Ded1 revealed that,

under saturating concentrations of Ded1, this substrate was unwound with a rate constant

similar to that for the chimeric substrate with 10 nt RNA at the 5’-end

130 Figure 6.3 A B 3’5’ 3’ 5’ * * * 3’5’ 3’ 5’ 30 * 6 10 3′ 5′ * 0.1 1 10 -1 k unwind [min ] 0 30 95 °C Time [min]

Figure 6.3 Ded1 unwinds duplexes lacking any free RNA terminus. Gray lines; DNA; black line, RNA; asterisks, the radiolabel. 5’ and 3’ represent the 5’-end and 3’-end of the respective strand. (A) Unwinding of a multi-piece substrate as depicted on top with 600 nM Ded1 and 2 mM ATP. The cartoons on the left

indicate mobility of single strands and duplexes. Number 6, 10, and 30 represent the length of the labeled

chimeric piece and DNA piece. Zero presents the reaction prior to the ATP addition. After the addition of

ATP, aliquots were removed from 30 s to 30 min. 95 °C represents the boiled reaction control. (B)

Unwinding rate constants (sliding points) of the MPS and the RNA-DNA chimeric substrate. Unwinding

rate constants were determined as in Fig.6.1. Ded1 was present at saturating concentrations.

131 (Fig.6.2A, Fig.6.4A). These data indicate that Ded1 can unwind substrates with RNA at

internal positions, suggesting that Ded1 can initiate unwinding within the helical region.

Removal of the overhang from the chimeric substrate prevented the fast

unwinding (Fig 6.4B). Unwinding reactions with increasing concentrations of Ded1 revealed that this substrate could not be saturated at experimentally accessible Ded1

concentrations (Fig.6.4A,B), suggesting that the overhang could aid the binding of Ded1

to the helical region lacking a RNA terminus. This substrate was unwound with a similar

rate seen for 16 bp RNA/DNA blunt-end substrate (compare Fig.6.4B and Fig.5.4D of

Chapter 5). This finding argues against the idea that 3 bp DNA at each end of the helical

region has to be frayed prior to strand separation. Such fraying would give rise a 10 bp

RNA/DNA duplex, which would be unwound much faster than the 16 bp RNA/DNA duplexes (cf. Chapter 3). Taken together, these results further support the notion that

Ded1 can initiate unwinding within the helical region.

To confirm and extend these observations, I designed a top strand with 5 nt DNA

at the 5’-end and 6 nt DNA at the 3’- end of 5 nt RNA and annealed this chimeric strand

to a 41 nt DNA bottom strand used before (Fig.6.4A, Fig 6.4C). The chimeric substrate

was clearly unwound by Ded1. However, at Ded1 concentrations of 1000 nM, the rate

constant for this substrate decreased almost 200-fold compared with the chimeric

substrate with 10 nt RNA within the helical region (compare Fig.6.4. A to C). Unwinding

reactions with increasing Ded1 concentrations revealed that this substrate could not be

saturated at experimentally accessible Ded1 concentrations, explaining the much reduced

unwinding rate constants for this substrate. Removal of the overhang from this substrate

did not decrease unwinding, contrary with the observations for the chimeric substrates

132 Figure 6.4

A E

DED1 ] MSS116

10 ] 5’ 3’ -1 4 -1 8 3310 * 3 6 2 4 2 1 Constant [min Constant Unwinding Rate Unwinding

* Constant [min 0

Unwinding Rate 0 0 200 400600800 0 100 200300400 035 s 0 min 035 s 0 min Time DED1[nM] Time MSS116[nM] ] B F ] 10 -1 -1 4 8 3 6 3310 * 2 4 2 1 * Constant [min Constant Constant [min Constant 0 Unwinding Rate

0 Unwinding Rate 035 s 0 min0 200 400600800 0 5 s 30 min 0 100 200300400 C Time DED1[nM] G Time MSS116[nM] ] ]

-1 1.0

-1 0.10 5’ 3’ * 0.08 0.8 655 0.06 0.6 0.04 0.4 0.02 0.2 Constant [min Constant Constant [min Constant

Unwinding Rate Rate Unwinding 0.0 * Unwinding Rate 0.00 0 200 400600800 0 100 200 300400 035 s 0 min 035 s 0 min DED1[nM] MSS116[nM] D Time H Time ] ]

-1 0.10 1.0 -1 0.08 0.8 655 * 0.06 0.6 0.04 0.4 * 0.02 0.2 Constant [min Constant Unwinding Rate Rate Unwinding Constant [min

035 s 0 minUnwinding Rate 0.00 035 s 0 min0.0 Time 0 200 400600800 Time 0 100 200 300400 DED1[nM] MSS116[nM]

Figure 6.4. Unwinding within the helical region by Ded1 and Mss116. Gray lines, DNA; black lines, RNA; asterisks, the radiolabel. The length of the RNA and DNA portion of the chimeric strand is indicated by the numbers underneath the cartoons on the left. 5’ and 3’ represent the 5’- and 3’-end of the DNA loading strand. Left panels: Unwinding by Ded1 (A, B, C, or D) or by Mss116 (E, F, G, or H). Right panels: Dependence of unwinding rate constants on Ded1 or Mss116 concentrations. For sequences, see table 5.1. (A) Left panel: Unwinding by 1000 nM Ded1. Right panel: The data were fitted to a sigmoidal isotherm,

-1 yielding k max = 7.8 ± 0.1 min , K d ′ = 598 ± 3 nM, and n = 15.3 ± 7.6. (B) Left panel: unwinding by 1000 nM Ded1. Right panel: Line represents a trend (C) Left panel: unwinding by 1000 nM Ded1. Right panel: Line represents a trend (D) Left panel: unwinding by 1000 nM Ded1. Right panel: Line represents a trend (E) Left panel: unwinding by 400 nM Mss116. Right panel: The data were fitted to a sigmoidal isotherm,

-1 yielding k max = 3.2 ± 0.1 min , K d ′ = 34 ± 3 nM, and n = 6.4 ± 1.6. Substrate is the same as in panel A. (F) Left panel: unwinding by 400 nM Mss116. Right panel: The data were fitted to a sigmoidal isotherm,

-1 yielding k max = 4.2 ± 0.1 min , K d ′ = 192 ± 5 nM, and n = 2.0 ± 0.1. Substrate is the same as in panel B. (G) Left panel: unwinding by 400 nM Mss116. Right panel: The data were fitted to a sigmoidal isotherm,

-1 yielding k max = 0.44 ± 0.01 min , K d ′ = 165 ± 3 nM, and n = 3.8 ± 0.2. Substrate is the same as in panel C. (H) Left panel: unwinding by 400 nM Mss116. Right panel: The data were fitted to a sigmoidal isotherm,,

-1 yielding k max = 0.43 ± 0.01 min , K d ′ = 146 ± 2 nM, and n = 3.6 ± 0.1. Substrate is the same as in panel D.

133 with 10 nt RNA, for which a clear decrease of unwinding rate constants was observed

after removal of the overhang (Fig.6.4B,D). Unwinding reactions of the substrate with 5

nt RNA lacking the overhang with increasing Ded1 concentrations revealed that this

substrate was unwound with a rate constant similar to that with the overhang at all Ded1 concentrations examined (Fig.6.4C,D). Thus, the stimulatory role of the overhang disappears when the RNA in the helical region is shortened beyond a critical length (5 nt). Nevertheless, the data clearly show that chimeric substrates with only a short RNA within the helical region can be unwound by Ded1. These observations provide strong support to the notion that Ded1 can be initiated from the internal positions.

6.1.5 Mss116 can also initiate unwinding within the helical region.

Next, I tested whether another DEAD-box helicase Mss116 can also initiate strand separation within the helical region. As Ded1, Mss116 readily unwound the chimeric substrate with 10 nt RNA within the helical region (Fig.6.4E), suggesting that

Mss116 can also initiate unwinding within the helical region.

At Mss116 concentrations of 400 nM, removal of the overhang from the chimeric

substrate did not decrease unwinding (Fig.6.4E, F). Unwinding reactions with increasing

Mss116 concentrations revealed that the overhang substrate was saturated at lower

Mss116 concentrations than the blunt-end substrate (Fig.6.4E, F). However, at saturating

Mss116 concentrations, both substrates were unwound with a similar rate constant. These data show that at low Mss116 concentrations, the overhang stimulates strand separation.

At saturating Mss116 concentrations, unwinding directly from the helical region is as fast as unwinding of the substrate with the overhang. These observations lend additional

134 support to the notion that the overhang does affect the actual strand separation by DEAD- box helicases.

To further confirm that Mss116 could unwind from internal positions, I tested the

ability of Mss116 to unwind the chimeric substrates containing 5 nt RNA. Mss116

separated this substrate readily (Fig.6.4G). At Mss116 concentrations of 400 nM,

removal of the overhang did not prevent fast unwinding (Fig.6.4H). Unwinding reactions

of the above substrates revealed that both substrates were saturated by Mss116 at similar concentrations. Therefore, the overhang does not aid the binding Mss116 to the helical region with 5 nt RNA. These observations parallel the findings with Ded1, where the loading function of the overhang disappeared when the RNA within the helical region was shortened to 5 nt. Both substrates were also unwound at similar rates at all Mss116 concentrations examined (Fig.6.4G,H), reinforcing the notion that the overhang did not have a role on actual strand separation. However, at saturating Mss116 concentrations, unwinding rate constants for these substrates were 7-fold lower than the substrate with 10 nt RNA(Fig.6.4G,H), indicating that the extent of RNA in the duplex region affects the rate constant of the strand separation process. Despite these rather unexpected effects, the data above clearly show that both Ded1 and Mss116 can unwind substrates with RNA at internal positions. As Ded1 and Mss116 are phylogenetically distant within the DEAD- box helicase family, our data suggest the ability to initiate unwinding within the helical region might be widespread among DEAD-box helicases.

6.1.6 Both Ded1 and Mss116 do not preferentially initiate unwinding from the

terminus of the helical region.

135 Having established that Ded1 and Mss116 could initiate unwinding from both

ends of the duplex and within the helical region, it was important to test whether Ded1 and/or Mss116 displayed a preference for termini over internal positions of the duplex.

Inspection of the unwinding rate constants of substrates with 10 nt RNA at either 5’, 3’ or

internal positions revealed that Ded1, at saturating concentrations, unwound substrates

containing RNA at the 5’ and internal positions with equal rate constants, whereas the

substrate with RNA at the 3’ position was unwound slower by a factor of 13. To test

whether preferential unwinding of substrates with RNA at the 5’- and internal positions

over at the 3’-position still holds when the overhang had no stimulatory role on

unwinding , I designed three 16 nt RNA/DNA chimeric strands with 5 nt RNA either at

the 5’-, 3’, or internal positions and annealed these strands to a 41 nt DNA piece used

before (Fig 6.4A and Fig.6.5). Unwinding rate constants of above substrates were measured at increasing Ded1 concentrations. None of the substrates could be saturated by

Ded1 (Fig.6.4C and data not shown), regardless of the RNA position. At Ded1 concentrations of 1000 nM, the substrates with the RNA at the 5’- and internal positions were unwound with a similar rate constant, which was, however, at least 3-fold faster than that observed for the substrate with the RNA at the 3’-position (Fig.6.5). Removal of the overhang from the above substrates did not significantly change unwinding rate constants under all Ded1 concentrations examined. These observations confirm the finding above (Fig.6.4C,D) that the loading function of the overhang disappeared for substrates with 5 nt RNA, regardless of the position of the RNA. Nevertheless, these results show that Ded1 does not preferentially initiate unwinding from the RNA terminus.

If anything, initiating unwinding from the 3’-position of the top strand is least favorable.

136 Figure 6.5

DED1 MSS116 5’ 3’ 11 5 5’ 3’ 11 5 5’ 3’ 655 5’ 3’ 655 5’ 3’ 5 11 5’ 3’ 5 11 0.001 0.01 0.1 1 0.001 0.01 0.1 1 -1 -1 k unwind [min ] k unwind [min ]

Figure 6.5 Both Ded1 and Mss116 do not preferentially initiate unwinding from the terminus. Gray lines, DNA; black lines, RNA. The sequence of the substrates were the same as the substrate used in panel C, figure 6.4. Left panel: Unwinding by Ded1 (sliding points). No substrates could the saturated by Ded1. Unwinding rate constants was determined as in panel A, figure 6.4 at the Ded1 concentration of 1000 nM. Right panel: Unwinding by Mss116.Unwinding rate constants were determined as in panel E, figure 6.4 at saturating Mss116 concentrations.

137 Unwinding reactions of above substrates at increasing Mss116 concentrations

revealed that these substrates were saturated by Mss116 at similar concentrations (data

not shown). At saturating Mss116 concentrations, the substrate with the RNA at internal

position were unwound at least 4-fold faster than that with the RNA at the 5’-position and

40-fold faster than that with the RNA at the 3’-position (Fig.6.5). As with Ded1, removal of the overhang from the above substrates did not significantly change unwinding rate

constants under any of the Mss116 concentrations examined (Fig.6.5), supporting the

idea that the loading function of the overhang required a RNA in the helical region longer

than 5 nt. Collectively, these results show that both Mss116 and Ded1 do not preferentially initiate unwinding from the RNA terminus for substrates with 5 nt RNA.

Thus, unwinding pattern observed for substrates with 10 nt RNA still hold even when the

overhang does not help loading.

6.1.7 The unwinding efficiency of both Ded1 and Mss116 decreases with the length

of RNA stretches within the duplex region.

Recent structural studies of the DEAD-box helicases VASA and eIF4AIII with

bound RNA revealed that those enzymes bound the phosphodiester backbone of five

nucleotides (75, 77, 131). Four (VASA) and three (eIF4AIII) of these contacts involved

2’-OH groups. Given that only 5 RNA nucleotides were sufficient for unwinding

(Fig6.5), it was thus interesting to ask whether all contacts to 2’-OH seen in the crystal

structures were necessary for duplex unwinding. To address this question, I designed a

top strand with 7 nt DNA at each end of 2 nt RNA and annealed this chimeric strand to a

16 nt DNA piece used before (Fig.6.4B and Fig 6.6). This substrate, albeit slower than

138 the substrates with 5 nt RNA, could be unwound by Ded1 (Fig.6.6A). Unwinding reactions with increasing Ded1 concentrations revealed that the substrate could not be saturated by Ded1 (data not shown). At Ded1 concentrations of 1000 nM, the substrate was unwound with a rate constant 16-fold slower than the substrate with 5 nt RNA within the helical region and 128-fold slower than the substrate with 10 nt RNA within the helical region (Fig.6.6A). Mss116 could also unwind the substrate with 2 nt within the helical region (Fig.6.6B). At saturating Mss116 concentrations, this substrate was unwound with a rate constant 97-fold slower than that with 5 nt RNA within the helical region and 795-fold slower than that with 10 nt RNA within the helical region (Fig.6.6B).

These results indicate that contacts to 2’-OH are important for the efficiency of strand separation but not essential for unwinding, as long as at least two 2’-OH groups are present.

139

Figure 6.6

AB16 16 n n ] ] -1 -1 DED1 MSS116 1.0 1.0 0.1 0.1 0.01 0.01 Constant [min Constant [min Constant Unwinding Rate Rate Unwinding Unwinding Rate 0.001 0.001 02 5 10 02 5 10

RNA length within RNA length within the helical region [nt] the helical region [nt]

Figure 6.6 The unwinding ability of both Ded1 and Mss116 decreases with the RNA length within the helical region. Gray lines, DNA; black lines, RNA; 16, 16 bp blunt-end duplex; n, the RNA length within the helical region. The sequence of substrates indicated by the cartoon on the top is the same as that in panel B, figure 6.4. (A) The dependence of unwinding rate constants of Ded1 on the RNA length within the duplex region. The unwinding rate constants were determined as panel B, Fig.6.4 at the Ded1 concentration of 1000 nM. Note that Ded1 cannot unwind the DNA/DNA substrates. (B) The dependence of unwinding rate constants of Mss116 on the RNA length within the duplex region. The unwinding rate constants were determined as panel F, Fig.6.4 at the Mss116 concentration of 400 nM. Note that Mss116 cannot unwind the DNA/DNA substrates.

140 6.2 Discussion

Here, I have shown that the length of single strand overhang affects the functional

affinity of Ded1 for duplex RNA but not the unwinding rate constants when Ded1 is

present at saturating concentrations. Under saturating Mss116 concentrations, the blunt-

end substrates are unwound with a rate constant similar to the substrates with an

overhang. I further show that Ded1 and Mss116 can initiate unwinding from either side

as well as internal positions of the duplex. Finally, both Ded1 and Mss116 can unwind

RNA/DNA hybrid substrates containing as few as 2 nt RNA. These observations lent further support to our previous conclusion that DEAD-box helicases do not translocate on one of the duplex strands as the unwinding mode. Instead, these data suggest that Ded1 and Mss116 switch duplexes apart into single strands by binding directly on the duplex region.

As established earlier (Chapter 5), Ded1 and Mss116 use a single stranded RNA for facilitating the association of the enzymes with the helical region (step 1, Fig.6.7, cf.

Chapter 5) but not for initiating translocation. Consistent with this observation, the data here indicate that the overhang does not have a role beyond loading the enzymes to the helical region. If the overhang could activate DEAD-box helicases, changing the overhang length should affect the unwinding rate constants. However, although the functional affinity for a substrate with 15 nt overhang is lower than for substrates with 25 nt and 35 nt overhangs, all of these substrates are unwound with a similar rate constant if

Ded1 is present at saturating concentrations (Fig.6.1). Moreover, Mss116 unwinds blunt-

141 Figure 6.7

Terminal Internal Initiation Initiation

DEAD-box protein binding facilitated by single stranded regions

partial helix destabilization

dissociation of remaining basepairs affects unwinding rate with longer duplexes

Figure 6.7 DEAD-box helicases unwind RNA duplexes from both internal and terminal helical regions. The single strand overhang (not shown for simplicity purpose) facilitates the loading of DEAD-box helicases to both terminal and internal helical regions. Then DEAD-box helicases disrupt part of the helical region by coupling to ATP binding and hydrolysis. Spontaneous dissociation of remaining basepairs leads to complete unwinding of the helical region.

142 end substrates with the same rate constant as substrates with an overhang as long as the

enzyme is presented at saturating concentrations (Fig.6.4, 6.5). These data indicate that

the single stranded overhang does not activate the actual strand separation by Ded1 and

Mss116. Our data now clearly and quantitatively show that the overhang loads DEAD-

box proteins on the helical region where unwinding is directly initiated.

Unexpectedly, the overhang does not assist the binding of either Ded1 or Mss116

to the helical region when the RNA length in the helical region decreases to 5 nt

(Fig.6.4C,D,G,and H). The functional affinity of Mss116 for the substrates with 5 nt

RNA is similar to the blunt-end substrate with 10 nt RNA (Fig.6.4E,F,G,and H). These

data suggest the enzymes loaded by the overhang sense the extent of RNA structures in

the helical region, whereas the enzymes that bind directly to the helical region do not.

After association of the enzymes, part of the duplex region is destabilized in an-

ATP-dependent manner (step 2, Fig.6.7). Structural studies of DEAD-box helicases bound to nonhydrolysable ATP analog AMPPNP and RNA reveal that double stranded

RNA has to be partially unwound in order to model it on the RNA binding site of the enzymes (75, 77, 131). However, I did not observe any strand separation by Ded1 and

Mss116 in the presence of AMPPNP, which mimics the ATP binding state (7, 9). If ATP binding is not sufficient for strand separation, strand separation should be coupled with other steps of the ATP hydrolysis cycle, i.e., hydrolysis of ATP to ADP·Pi, or release of

ADP or Pi. Another challenging question is the coupling efficiency between the ATP hydrolysis and strand separation, i.e., how many ATPs are hydrolyzed before a strand separation event occurs?

143 After active strand separation, the remaining basepairs most likely dissociate

spontaneously (step 3, Fig.6.7). Although Ded1 and Mss116 cannot unwind DNA/DNA

duplexes, both enzymes can unwind RNA/DNA chimeric duplexes where part of the duplexes are substituted with DNA/DNA regions (Fig.6.2 and Fig 6.4). Therefore, upon

opening a region at RNA/DNA duplex, the DNA/DNA duplex should dissociate

spontaneously, leading to complete strand separation. In addition, if strand separation

involves a step of spontaneous strand dissociation, the unwinding reactions should be

sensitive to the duplex stability. Indeed, unwinding rate constants of both Ded1 and

Mss116 scale with the length and stability of RNA duplexes (Chapter 3 and ref. (7)).

Moreover, unwinding reactions of Ded1 and Mss116 are highly temperature sensitive

(data not shown), suggesting that unwinding reactions may involve a step with a high

activation energy, which points to a rate limiting process depending on spontaneous

dissociation of RNA structure.

Notably, if the unwinding reactions are initiated from internal positions, the

remaining basepairs will be located at either side of the actively destabilized region and

therefore less stable than the remaining basepairs for the reactions initiated from the

terminus, where the remaining basepairs are located at one side of the actively

destabilized region (step 3, Fig.6.7). If the rate constant of the dissociation of the

remaining basepairs is comparable to the rate constants of other unwinding steps, these

unwinding reactions initiated within the duplex should be faster than those initiated from

the terminus. Consistent with this notion, under saturating Mss116 concentrations where

association of Mss116 with the helical region is not rate limiting, the chimeric substrates

144 with 5 nt RNA in internal positions were unwound at least 4-fold and 40-folder faster than those with RNA at 5’-end and at 3’-end, respectively (Fig.6.5).

Collectively, these data suggest that the stability of the duplex influences

unwinding rate constants twice. First, the fraying of the helical region prior to or during enzyme binding, which depends on local stabilities. Second, upon ATP-hydrolysis driven destabilization of the duplex, the dissociation of remaining basepairs. Consistent with this notion, unwinding rate constants of Ded1 do not correlate linearly with the overall stability but are affected strongly by the local stability of the duplex as well (unpublished data). Nonetheless, it is unclear how the local stability and overall stability of the duplex quantitatively affect unwinding rate constants. Our model proposed here provides a possible guiding principle to dissect the influence of each step on unwinding reactions. In the future, it is crucial to understand how DEAD-box helicases exploit these mechanistic features for their physiological functions.

145 Chapter 7: Future Directions

In this thesis, I have demonstrated that the DEAD-box RNA helicases Ded1 and

Mss116 unwind duplexes in a manner that significantly differs from helicases that

translocate unidirectionally along single stranded RNAs/DNAs. Ded1 and Mss116 do not

utilize a single stranded region to initiate translocation but to attain close proximity to the

duplex, where strand separation is directly initiated (Chapter 5). I further show that Ded1

and Mss116 can initiate unwinding from either terminus as well as from internal

positions of the duplex, lending further support to the notion that DEAD-box helicases do

not translocate on one of the duplex strands for strand separation (Chapter 6). The new

type of helicase mode shown here for Ded1 and Mss116 explains observations with other

DEAD-box proteins and is consistent with the biological activities of DEAD-box

helicases, the strict local remodeling of RNA-protein and RNA-RNA complexes (Chapter

3, 4).

Data I have collected so far suggest an unwinding mechanism that involves the steps shown below.

Spontaneous Active dissociation Complete Fraying strand Strand Loading of remaining separation basepairs separation + ATP

ct ire ng D di k in d Fraying b

146 The next step in elucidating the mechanism of duplex unwinding by DEAD-box

proteins is the systematic and quantitative verification of this reaction scheme. In the

following paragraphs, I outline possible strategies to test this hypothetical mechanism.

7.1 Binding of Ded1 and Mss116 to the helical region

Unwinding of duplex RNA by Ded1 and Mss116 is promoted by an adjacent

ssRNA, which does not have to be connected to duplex RNA. However, Ded1 and

Mss116 do not unwind dsRNA by translocating from ssRNA, suggesting that the

association of Ded1 and Mss116 with ssRNA facilitates the binding of the enzymes to the

adjacent dsRNA. The complex of DED1 and RNA in the absence of ATP is not stable

(dissociation rate constant greater than 5 min-1, unpublished data). Tight binding of Ded1

occurs in the presence of the nonhydrolyzable ATP analog, AMPPNP, suggesting that

ATP binding increases the affinity of Ded1 for RNA. However, binding of Mss116 to

RNA is not enhanced by ATP binding.

Two questions about the association of enzymes with the helical region remain

unanswered. First, does the enzyme unit on the single strand have to transfer to the

helical region without leaving the nucleic acid or alternatively, does the enzyme unit

recruit another functional enzyme unit to the helix in order to unwind the duplex? I

believe the latter model is more likely because: 1) AMPPNP does not inhibit unwinding

rate constants of Ded1 when the AMPPNP concentration is below 100 μM, whereas

inhibition of the ATPase activity with this AMPPNP concentration is profound

(unpublished data). If there is only one functional DED1 unit, one would expect that

unwinding and ATPase activities of DED1 would be equally sensitive to the AMPPNP

147 inhibition. It is conceivable that, during the unwinding reactions, there are at least two

functional DED1 units, which have distinct affinities for AMPPNP. It is very likely that

Ded1 that binds an overhang has a higher affinity for AMPPNP than that binds blunt-end

duplex. To test this hypothesis, it is necessary to measure Ded1’s ATPase activities with increasing concentrations of AMPPNP in the presence of either single stranded RNA or

blunt-end duplex. The ATPase activity of Ded1 with ssRNA should be more sensitive to

AMPPNP inhibition than that with blunt-end duplex. 2) I have found that Ded1 binds an

overhang tightly with a slow dissociation rate constant (lesser than 1 × 10-3 min-1) in the

presence of 0.1 mM AMPPNP. The unwinding rate constant under the same AMPPNP concentration is greater than 3 min-1. Therefore, it is unlikely that the duplex is unwound by the Ded1 unit bound to the overhang. Ded1 should remain binding the overhang during the unwinding reactions. To test this idea, unwinding reactions with a 16 bp RNA duplex with a 25 nt overhang in the presence of 0.1 mM AMPPNP should be performed and simultaneously the binding of Ded1 to the loading strand and top strand on native gels should be examined. The loading strand should still be bound by Ded1, while the unwound top strand should not.

The second question asks whether the helical region has to be frayed prior to

binding of the enzyme? Structural studies reveal that the helical region has to be partially unwound to accommodate an A-form RNA helices to RNA binding site of DEAD-box

proteins (75, 77, 131), suggesting that duplex RNA may have to be frayed prior to the association of the enzymes. If this is true, Ded1 will preferentially bind to the least stable region of duplex RNA. To test this possibility, the binding of Ded1 to the helical region by nuclease protection assay could be probed.

148

7.2 Part of the RNA duplex is actively unwound by DED1 in an ATP-dependent

manner.

I have shown that both Ded1 and Mss116 can unwind a duplex with both ends

blocked by a DNA region, suggesting that unwinding can be initiated from the internal

positions of the helical region. To provide more direct evidence for this new type of

strand separation mode, it would be advantageous to use DNA footprinting to detect

which of the basepairs inside the duplex are opened while the basepairs at the ends are

closed.

Next, it would be important to investigate which step of the ATP hydrolysis cycle

is coupled with strand separation. ATP binding is not sufficient for stand separation, as

both Ded1 and Mss116 can not unwind substrates in the presence of AMPPNP. However,

Ded1 can separate 13 bp but not 16 bp duplex RNA in the presence of ADP·AlF4, an

ATP analog mimicking the transition state of ATP hydrolysis, suggesting that the actual

cleavage of ATP is not necessarily required for strand separation for at least very short duplexes (Liu and Jankowsky, unpublished data). Therefore, strand separation could be coupled to either the transition state of ATP hydrolysis or the phosphate/ADP release from the post hydrolysis state of ADP·Pi. To test these two possibilities, it will be interesting to examine duplex unwinding in the presence of ADP·BeF(x), an ATP analog

mimicking the post hydrolysis state of ATP.

The most critical and also most challenging question is how many ATPs are

required for each strand separation event. To answer this question, it is necessary to

determine how many ATP hydrolysis events occur prior to a strand separation event. It

149 may be possible to directly address this question with a single molecule FRET system

where binding of ATP to Ded1 and strand separation events are monitored

simultaneously.

7.3 After the active helix opening, remaining basepairs dissociate spontaneously.

I have found that the unwinding rate constants scale with the length and stability

of the duplex. Moreover, unwinding rate constants are highly temperature sensitive.

Preliminary data suggest an activation energy greater than 160 kJ/M, pointing toward a

process involving spontaneous strand separation. To confirm that the spontaneous

dissociation of remaining basepairs is indeed the process causing this high activation

energy, it will be useful to systematically measure the temperature sensitivity of

unwinding reactions using duplexes with different lengths. As the duplex shortens, the

number of remaining basepairs after the active duplex separation will decrease, which

should decrease the activation energy of unwinding reactions.

Before the spontaneous dissociation of remaining basepairs, DED1 can fall off the duplex, which allows reannealing of opened basepairs. I believe this step contributes

significantly to the observed reaction amplitude under single cycle conditions. If this is true, I expect that unwinding amplitudes under single-cycle conditions will scale with the stability of the duplexes. As the stability of the duplex decreases, the reaction amplitudes should increase.

150 Chapter 8: Materials and Methods

Ded1 expression and purification.

To facilitate the purification of Ded1, 18 nucleotides that encodes hexahistidine amino acid residues was added to N-terminus of Ded1 cDNA. This cDNA was cloned into a pET22b plasmid. The resulting plasmid was transformed into the E. coli strain DE3

(BL21), a strain deficient in RNaseE. Cells were grown in LB media (10 g Bacto- tryptone, 5 g yeast extract, and 5 g NaCl) at 37 ºC by vigorously shaking (~250 rpm). The cells were grown in LB media containing 100 µg/ml Ampilicin to preserve the Ded1- containing plasmid in cells. When the absorbance at 600 nm (A600) reached ~0.6, the expression of Ded1 was induced by addition of 0.5 mM IPTG (final concentration) and the induction was continued overnight at 28 ºC. Higher temperatures caused the formation of insoluble inclusion bodies of Ded1. Cells were harvested by centrifugation

~5000 rpm for 30 min. The pelleted cells were suspended in buffer A (50 mM NaH2PO4, pH 6.0, 300 mM NaCl, 10 % glycerol with freshly prepared 5 mM imidazole, 0.5 %

NP40, 0.2 mM PMSF and 1 pellet of CompleteTM protease inhibitor cocktail tablets

(Roche)). Cells were broken by sonicating cells 10 times (sonic dismembrator model 100,

Fisher Scientific) at output power 10 W on ice, with a 1 min interval to prevent cells from overheating. After lysis, centrifuge for 30 min at 15, 000 rpm in a compact microcentrifuge (Eppendorf 5415) to get rid of the particles and debris and the soluble fraction containing Ded1 was collected and incubated with Ni-NTA agarose beads

(Qiagen) overnight. Beads were loaded into a 10 cm x 1 cm (length x diameter) column

151 and were washed extensively in sequential manner with 20 mL buffer A with 5 mM imidazole and 700 mM NaCl, with 30 mM imidazole, and with 60 mM imidazole before elution with 250 mM imidazole. The high salt in the first washing step is employed to get rid of the RNase that otherwise will be enriched by Ni-NTA beads. Fractions containing

Ded1 were diluted six times in buffer B (50 mM Tris-HCl (pH 8.0), 2 mM DTT, 1 mM

EDTA, 10 % Glycerol, 0.1 % Triton-X 100) in order to absorb Ded1 to phosphocellulose

(P11, Whatman). Ded1 was eluted from the phosphocellulose with 300 mM NaCl in buffer B after washing twice with 100 mM NaCl and 200 mM NaCl in buffer B.

Homogeneity (>98%) of Ded1 was assessed by SDS PAGE and Coomassie staining. The concentration of Ded1 was determined with Bradford method and bovine serum albumin

(BSA, New England Biolabs) was used to obtain the standard curve of A595nm vs. the

BSA concentrations for calibration. The Ded1ΔC mutant, in which 83 amino acid residues at the carboxyl-terminus of Ded1 were deleted, was generated by replacing the codon of

T519 with a UAA stop codon using PCR based mutagenesis. The PCR products were cloned into pET-22b and sequenced. The resulting plasmid was then transformed into

BL21(DE3). Ded1ΔC was expressed and purified as wild type Ded1. Homogeneity

(>90%) and the concentration of Ded1ΔC was assessed by SDS PAGE and Coomassie staining. NPH-II was expressed in baculovirus infected insect cells and purified as described (25). Purified eIF4A was a gift from Dr. W. Merrick (158).

152 RNA substrates preparation.

RNA oligonucleotides were purchased from DHARMACON and DNA

oligonucleotides were purchased from Sigma-Genosys. Single stranded RNAs/DNAs

(ssRNA/DNAs) were labeled with [γ32P-ATP] by T4 polynucleotide kinase (T4 PNK)

(62). In general, reaction mixture (10 μL) containing 100 pmoles ssRNA/DNAs, 100 pmoles unlabeled ATP, 50 pmoles [γ32P-ATP], and 1x T4 PNK buffer were incubated

with 1.5 u/μL T4 PNK enzyme for 1 h at 37 ºC. The full length labeled ssRNA/DNAs

were separated from degraded nucleic acid strands on a 20 % denaturing polyacrylamide

gels with acrylamide:bis ratio of 29:1 in a buffer containing 7 M urea and TBE buffer (89

mM Tris-HCl (pH8.0), 89 mM Borate, and 2 mM EDTA) at 40 V/cm. Then cut out the

gel pieces containing the full length RNA/DNAs and crush the gels through a syringe into

1.6 mL eppendorf tube. Add 600 μL elution buffer to the tube and elute the RNA/DNAs overnight at cold room by rotating slowly on a rotator (Bardstead). Draw off elution buffer from gel slice, spin down remaining acrylamide, and decant buffer. Split elution buffer into two 1.6 mL eppendorf tubes. Add three times volumes of 100% ethanol (~900

µL) and 1μL glycerol to each tube. Precipitate the oligos on dry ice for 30 min. Pellet the oligos by centrifugation at 13, 000 rpm for 30 min at cold room in a compact microcentrifuge (Eppendorf 5415). Decant supernatant and drip dry on a tissue paper.

Dry the oligos in a speedvac concentrator (SVC100H, Savant) connected to a refrigerated vapor trap (Thermo Electron Corporation) for 15 min. Dissolved the oligos in 17 μL water. Duplex RNA/DNA duplexes were formed by annealing the labeled ssRNA/DNAs with 100 pmoles unlabeled complementary ssRNA/DNAs in annealing buffer (10mM

153 MOPS (pH 6.5), 1 mM EDTA, 50 mM KCl) by heating the reactions to 95 ºC for 2 min, followed by cooling down at room temperature slowly over 2 h. The duplex substrates were separated from ssRNA/DNAs on 15% non-denaturing polyacrylamide gels with acrylamide:bis ratio of 29:1 in 0.5 time TBE buffer (pH8.0) at 20 V/cm and were purified as described above for the single stranded oligos. The concentration of the labeled ssRNA/DNAs and duplex substrates were quantified by a liquid scintillation counter (62).

ATPase reactions

ATPase reactions were performed as described (250). Briefly, reaction mixtures

(10 μL) containing 40 mM Tris-HCl (pH 8.0), 20 mM NaCl, and 0.01% NP40 were

incubated with 150 nM Ded1 and, where present, 1 μM ssRNAs or 500 nM duplex RNA substrates at room temperature for 5 min. Longer incubation did not alter the observed rate of ATP hydrolysis. ATPase reactions were initiated by adding an equal molar mixture of 0.5 mM ATP/MgCl2. The ATP mixture contained a trace amount of [γ32P-

ATP] with a ratio of 1000 to 1 in order to indicate the hydrolysis of ATP. At the times indicated, aliquots (1 μl) were removed and spotted on a polyethyleneimine cellulose thin layer chromatography (PEI TLC) plates (Fisher Scientific). The free inorganic [32P] phosphates were separated from [γ32P-ATP] in the TLC buffer containing 1 M formic

acid and 0.75 M LiCl for 20 min. Subsequently, the TLC plates were drip-dried in a

laboratory fume hood. The radiolabeled [32P] and [γ32P-ATP] were visualized with a

Molecular Dynamics Phosphorimager and the ImageQuant 5.2. software (Molecular

Dynamics) (25).

154

Equilibrium binding

Reaction mixtures (20 μL) containing 40 mM TrisHCl (pH 8.0), 50 mM NaCl, 0.5 mM MgCl , 2 mM DTT, 1 unit/μL RNasin, 0.01% NP40, and 0.5 nM radiolabeled RNA 2 substrate were incubated with Ded1 and, where present, with 1 mM AMPPNP (+1 mM

MgCl ) for 5 min in a temperature-controlled aluminum block at 19°C. The Ded1 2 concentration was 600 nM if not explicitly indicated. Longer incubation times did not alter the observed binding behavior. Subsequently, 2 μM (final) single stranded RNA (73 nucleotides) and glycerol (50% v/v, final) were added to each sample. The single stranded RNA sequestered excess and non-stably bound Ded1, which left only stable

Ded1-RNA complexes. To measure the dissociation rate constant of Ded1 from the RNA substrates, each sample was further incubated with the same 2 μM ssRNA for the indicated times. The unlabeled ssRNA prevented the rebinding of Ded1 to the RNA substrates. Samples were then immediately applied to non-denaturing 8 % polyacrylamide gels with acrylamide:bis ratio of 37.5:1 in a Tris-Glycine buffer (25 mM

Tris-HCl (pH8.3), 192 mM glycine) without SDS. Ded1-RNA complexes were separated from free RNA at 4°C at 20 V/cm. Subsequently, gels were dried, and the radiolabeled

RNAs were visualized and quantified with a Molecular Dynamics Phosphorimager and the ImageQuant 5.2. software (Molecular Dynamics).

155 Limited alkaline hydrolysis of ssRNA oligonucleotides

Reaction mixture (4 μL) containing 6 M urea and 50 mM NaOH was incubated

with 2 μL 200 μM 32P-labeled RNA oligonucleotide (5’-UUAGUACGUCCCAGACAG

CAUUGUACCCAGAGUCUGUACGG-3’) for 4 min in a temperature controlled

aluminum block at 85°C. The hydrolysis reactions were quenched by addition of 200 μL

quenching buffer (1 mM HCl and 10 mM Tris-HCl, pH 8.0). Then the hydrolyzed RNA

oligos were ethanol precipitated as described above and dissolved in 20 μL RNase-free

water.

Unwinding reactions

Reaction mixtures (30 μl) containing 40 mM Tris-HCl (pH 8.0), 50 mM NaCl, 0.5

32 mM MgCl2, 2 mM DTT, 1 unit/μl RNasin, 0.01 % NP40, and 0.5 nM [ P]-labeled RNA

substrate were incubated with Ded1 for 5 min. The Ded1 concentration was 600 nM

where not explicitly indicated. Reactions were performed in a temperature controlled

aluminum block at 19°C. All reactants were incubated at 19°C prior to the reaction to

avoid temperature changes upon addition of components. Unwinding reactions were

initiated by adding an equimolar mixture of MgCl2 and ATP to the indicated ATP

concentrations. For reactions including ADP and AMPPNP, ADP-MgCl2 and AMPPNP-

MgCl2 were added together with ATP-MgCl2 at the reaction start. At the times indicated,

aliquots (3 μl) were removed and the reaction was stopped with a buffer containing 1%

SDS, 50 mM EDTA, 0.1 % xylene cyanol, 0.1% bromophenol blue, and 20% glycerol.

156 Aliquots were applied to 15% non-denaturing polyacrylamide gels with acrylamide and bis ratio of 29 to 1 and duplex and single stranded RNA were separated at room temperature at 20V/cm. Subsequently, gels were dried and the radiolabeled RNAs were visualized and quantified with a Molecular Dynamics Phosphorimager and the

ImageQuant 5.2. software (Molecular Dynamics).

Annealing reactions

Reaction mixtures (30 μl) containing 40 mM Tris-HCl (pH 8.0), 50 mM NaCl, 0.5 mM MgCl2, 2 mM DTT 1 unit/μl RNasin, 0.01 % NP40 were incubated with Ded1 and,

where indicated, with MgATP, MgADP, or MgAMPPNP for 5 min. The Ded1

concentration was 600 nM or indicated explicitly otherwise. The duplex RNA substrates

were denatured at 95 ºC for 2 min and snap cool on ice for 1 min to prevent the

spontaneous reannealing of denatured single strand RNAs. Reactions were performed in a

temperature controlled aluminum block at 19°C. Annealing reactions were initiated by

addition of 0.5 nM of the denatured substrate strands. Aliquots (3 μl) were removed at the

times indicated and reactions were stopped with the same buffer used to quench the

unwinding reactions. Single stranded and duplex RNAs were separated as described for

the unwinding assays.

157 Thermal melting curves

Melting temperatures of complexes A and B (cf. Chapter 4, table 4.1) were determined in reaction buffer containing 40 mM Tris-HCl (pH 8.0), 50 mM NaCl, 0.5 mM MgCl2, 2 mM DTT, 1 unit/µL RNasin, 0.01% NP40. Complexes (A or B) at 0.5 nM were incubated at indicated temperatures ranging from 24 to 95 ºC in a temperature calibrated PCR machine for 5 min. Then reactions transferred on ice and buffer containing 1% SDS, 50 mM EDTA, 0.1% xylene cyanol, 0.1% bromophenol blue, 20% glycerol was added. Subsequently, aliquots were applied to a 15% non-denaturing polyacrylamide gels, duplex and single-stranded RNAs were separated, and the fraction of single stranded RNAs were determined and plotted vs. incubation temperature.

Structure conversion reactions

Reactions were performed at 24 ºC in a volume of 30 µL in a buffer containing 40 mM Tris-HCl (pH 8.0), 50 mM NaCl, 0.5 mM MgCl2, 2 mM DTT, 1 unit/µL RNasin,

0.01% NP40. For the conversion of structure A into B, 2 nM pre-formed complex A and

8 nM of strand x (final concentrations) were used; for the conversion of structure B into

A, 2 nM pre-formed complex B, and 6 nM strand x and 2 nM strand z were used, in order to maintain equal concentrations of all individual strands (x,y,z) in all reactions. Complex and single stranded RNAs at these concentrations were incubated for 5 min at reaction temperature. The reactions were initiated by addition of 800 nM Ded1 and an equimolar mixture of ATP and MgCl2 at the concentrations indicated. To the reactions without

158 Ded1, protein storage buffer (50 mM Tris-HCl (pH 8.0), 2 mM DTT, 1 mM EDTA, 10 %

Glycerol, 0.1 % Triton-X 100, and 300 mM NaCl) was added. Where applicable, 120 nM

U1A was added. At the times indicated, aliquots (3 µl) were removed and reactions were terminated on ice with 3 µl of stop buffer containing 1% SDS, 50 mM EDTA, 0.1% xylene cyanol, 0.1% bromophenol blue, and 20% glycerol. Subsequently aliquots were applied to non-denaturing polyacrylamide gels. Gels were dried and bands corresponding to single stranded and complex RNAs were visualized and quantified with a

Phosphorimager (Molecular Dynamics).

The distribution of the complexes A and B at the thermodynamically dictated

equilibrium was determined by incubating either 2 nM pre-formed complex A and 8 nM

of strand x (final concentrations), and, in an alternative reaction, 2 nM pre-formed

complex B, and 6 nM strand x and 2 nM strand z at 95 for 2 min in reaction buffer and

subsequent cooling of the respective mixture to reaction temperature over 2 h. The

mixture was then treated as the reaction aliquots above, i.e., placed on ice, combined with

stop buffer and applied to non-denaturing polyacrylamide gels.

Ensemble fluorescence measurements

In a Fluoromax 2 fluorometer (Spex), reaction mixtures (100 µL) containing 10

nM Cy5-labeled strand x and 10 nM Cy3-labeled complex A (10 nM) was incubated in

the reaction buffer (40 mM Tris-HCl (pH 8.0), 50 mM NaCl, 0.5 mM MgCl2, 2 mM

DTT, 1 unit/µL RNasin, 0.01% NP40) for 5 min at 24 ºC. Increasing incubation time up

to 30 min did not alter the results. Where applicable, 800 nM Ded1 and/or an equimolar

159 mixture of ATP and MgCl2 (at the concentrations indicated) was included. Then reaction

mixture was excited at a wavelength of 515 nm (with a bandwidth of 5 nm). Fluorescence

emission was measured from 530 to 750 nm (with a bandwidth of 5 nm).

Single molecule fluorescence measurements

Single molecule fluorescence was measured in a home-built total internal

reflection (TIR) microscope setup. The system is analogous to that described by Zhuang

et al. (228). Core components include a Nikon TE300 microscope, Pentamax Gen III

CCD (Princeton Instruments), and a 50 mW, doubled solid state laser

(Crystalser). Samples (50 pM Cy3 labeled complex A) were immobilized in a flow cell

coated with polyethyleneglycol (PEG) to prevent non-specific protein absorption (226,

251). Briefly, a mixture of PEG-NHS (MW 3000 -5000) and biotinylated PEG-NHS

(MW 3000) (SHEARWATER) was covalently attached to the flow cells that were amino- functionalized with Vectabond (VECTOR). Biotinylated samples were immobilized through streptavidin (MOLECULAR PROBES) to the biotinylated PEG (226, 251). Non- specific binding of the labeled RNA complex, which was tested by omission of streptavidin, was found to be less than 5% of the specific binding. Molecules were immobilized at an average distance of at least 2-3 µm between individuals, i.e., at multiples of the diffraction limit. Photobleaching studies confirmed that more than 90% of the fluorescent spots corresponded to single fluorophors. Control reactions measuring

Ded1-RNA binding verified that inactivation of Ded1 under the conditions of the single molecule experiments was insignificant (data not shown).

160 Reactions were conducted in the buffer used for the above experiments, except that 5 % (v/v) glucose were added together with an oxygen scavenging system consisting of glucose oxidase (SIGMA) and catalase (SIGMA) (228). Control reactions confirmed that these conditions did not significantly affect the reaction kinetics in solution (data not shown). Samples were excited by TIR (532 nm) and the fluorescence was collected through a 60 x 1.2 N.A. water immersion objective (NIKON). Excitation light was blocked with a 550 nm long path filter (OMEGA). Donor and acceptor fluorescence images were split by a diachronic filter (650 nm, OMEGA) and the two resulting paths were projected on one half of the intensified CCD. Frames were collected at a rate of 10 fps using customized software (228). Fluorescence and corresponding FRET values were computed as described (228). Data analysis was conducted with customized Matlab routines (228).

Data analysis for unwinding and annealing reactions

The fraction of single stranded and duplexed RNAs were determined from the relative amount of radioactivity in the respective bands (Iss, Idpx), measured with the

ImageQuant 5.2 software (Molecular Dynamics). The fraction of single stranded RNA was determined according to:

Frac ss = Iss·(Iss + Idpx)-1 (eq. I).

The fraction of duplex RNA was computed according to

Frac dpx = 1- Frac ss (eq. II).

161 (unw) Observed rate constants kobs for unwinding reactions were determined by fitting time courses to the integrated rate law for a homogenous first order reaction:

Frac ss = A·(1-e(-kobs(unw)·t)) (eq. III).

A is the reaction amplitude. Fitting the data to other rate equations yields curves

(ann) significantly deviated from the data points. Observed rate constants kobs for annealing

reactions were determined by fitting time courses to the integrated rate law for a homogenous first order reaction according to:

Frac ss = A·e(-kobs(ann)·t) (eq. IV).

(A is the reaction amplitude). Note that the annealing rate constants throughout

this paper are given as observed first order rate constants, valid at the fixed concentration

of Ded1 (600 nM), and RNA (0.5 nM for each strand or the duplex). Rate constants for

unwinding (kunw) and annealing (kann) under the ATP-dependent steady state between

duplex formation and strand separation were determined based on the following kinetic

scheme for a steady state between single stranded (ss) and duplex (dpx) RNA:

d[ss]/dt = kunw[dpx] – kann[ss] (eq. V)

Solving the differential equation, integrating, and rearranging for the fraction of single stranded RNA yields:

-1 -(kunw+kann)·t Frac ss = kunw·(kunw + kann) ·(1-e ) (eq. VI)

Eq. VI was used to fit the time courses of unwinding reactions at varying ATP,

ADP and AMPNP concentrations. The dependencies of rate constants for unwinding and

or annealing or ratios of rate constants on ATP, ADP, or AMPPNP concentrations was

analyzed by either fitting the respective curves to a hyperbolic binding isotherm

max -1 kx = kx ·[NTP] ·([NTP] + Km) (eq. VII),

162 or to a sigmoidal Hill binding isotherm

max n n n -1 kx = kx ·[NTP] ·([NTP] + Km ) (eq. VIII).

(kx is the value for the rate constant at a given ATP, ADP or AMPPNP

max concentration, kx the rate constant (or ratio of rate constants) at ATP, ADP or

AMPPNP saturation, and Km represents the ATP, ADP or AMPPNP concentration where

the half maximal effect is reached, n is the Hill coefficient). Curve fitting was performed

with Kaleidagraph (Synergy software).

Data analysis for the equilibrium binding reactions

The fraction of free RNA substrates and Ded1 bound RNA complexes were

determined from the relative amount of radioactivity in the respective bands (Ifree, Ibound).

Fraction bound was determined by according to:

Frac bound = Ibound·(Ifree + Ibound)-1 (eq. IX).

The dependency of fraction bound on the Ded1 concentrations was analyzed by

fitting the respective curves to a sigmoidal Hill binding isotherm assuming that all free

substrates were bound by Ded1 at enzyme saturation:

n n n -1 Frac bound = 1.0 ·[Ded1] · ([Ded1] + Kd ) (eq. X).

Kd represented the Ded1 concentration where half of the substrate was bound by

Ded1, n is the Hill coefficient.

The dissociation of Ded1 from the RNA substrate was analyzed by fitting the

respective curves to the integrated rate law of a homogenous first order reaction

according to:

163 (-koff . t) Frac bound = [Frac bound]0 · (e ) (eq. XI).

[Frac bound]0 represents the bound fraction prior to the addition of the RNA scavenger. koff represents the dissociation rate constant of Ded1 from the respective RNA

substrates.

Data analysis for the structure conversion.

The fraction of ssRNA Y and duplexed RNAs (YZ) and (YX) were determined

from the relative amount of radioactivity in the respective bands (IY, I(YZ), I(YX)),

measured with the ImageQuant 5.2 software (Molecular Dynamics). Fraction (YX) was

determined according to:

Frac (YX) = I(YX) ·( IY + I(YZ) + I(YX)-1 (eq. XIII).

Observed rate constants kobs for the formation of (YX) from (YZ) for the structure

conversion reactions started from (YZ) were determined by fitting time courses to the

integrated rate law for a homogenous first order reaction:

Frac (YX) = A·(1-e(-kobs· t)) (eq. XIV).

A is the reaction amplitude. Observed rate constants kobs for the disruption of

(YX) to form (YZ) for the structure conversion reactions started from (YX) were determined by fitting time courses to the integrated rate law for a homogenous first order reaction:

Frac (YX) = (1-A) · (e(-kobs· t)) + A (eq. XV).

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