Current Perspectives in microRNAs (miRNA)
Shao-Yao Ying Editor
Current Perspectives in microRNAs (miRNA)
Editor Shao-Yao Ying University of Southern California Los Angeles, CA USA
ISBN 978-1-4020-8532-1
e-ISBN 978-1-4020-8533-8
Library of Congress Control Number: 2008930759 © 2008 Springer Science + Business Media B.V. No part of this work may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording or otherwise, without written permission from the Publisher, with the exception of any material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Printed on acid-free paper 9 8 7 6 5 4 3 2 1 springer.com
Contents
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Structures of MicroRNA Precursors ..................................................... Piotr Kozlowski, Julia Starega-Roslan, Marta Legacz, Marcin Magnus, and Wlodzimierz J. Krzyzosiak
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Small RNA Technologies: siRNA, miRNA, antagomiR, Target Mimicry, miRNA Sponge and miRNA Profiling ...................... Guiliang Tang, Yu Xiang, Zhensheng Kang, Venugopal Mendu, Xiaoyun Jia, Qi-Jun Chen, Xiaohu Tang, and Xiaoqing Tang
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RNA Interference Expression Vectors Based on miRNAs and RNA Splicing ................................................................................ Akua N. Bonsra, Joshua Yonekubo, and Guangwei Du
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Recent Application of Intronic MicroRNA Agents in Cosmetics ......................................................................................... Shi-Lung Lin, David T.S. Wu, and Shao -Yao Ying
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MicroRNA Profiling in CNS Tissue Using Microarrays .................. Reuben Saba and Stephanie A. Booth
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MicroRNA and Erythroid Differentiation ......................................... Mei Zhan and Chao-Zhong Song
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Homeotic miRNAs: From Development to Pathologies ....................... Maya Ameyar-Zazoua, Irina Naguibneva, Linda Pritchard, and Annick Harel-Bellan
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MicroRNA in Muscle Development and Function ............................... Zhongliang Deng and Da-Zhi Wang
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MicroRNAs and Regenerative Medicine ............................................... Ji Wu and Zhaojuan Yang
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Role of mir-302 MicroRNA Family in Stem Cell Pluripotency and Renewal .................................................................... Shi-Lung Lin and Shao-Yao Ying
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Epigenetic Regulation of miRNA in Stem Cells ................................. Keith Szulwach, Xuekun Li, Xinyu Zhao, and Peng Jin
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Identification of Cellular Targets for Virally-Encoded miRNAs by Ectopic Expression and Gene Expression Profiling ...... Mark A. Samols, Rebecca L. Skalsky, and Rolf Renne
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MicroRNA in Neuropsychiatric Diseases ............................................ Evgeny I. Rogaev, Denis V. Islamgulov, and Anastasia.P. Grigorenko
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Role of Repeat-Associated MicroRNA(ramRNA) in Fragile X Syndrome (FXS) ............................................................... Shi-Lung Lin and Shao -Yao Ying
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miRNA and Schizophrenia ................................................................... Diana O. Perkins and Clark D. Jeffries
16 SNPs in microRNA and microRNA Target Sites Associated with Human Cancers ................................................... Shi-Hsiang Shen and Zhenbao Yu 17
Expression and Function of microRNAs in Chronic Myeloid Leukemia ................................................................................. Michaela Scherr, Letizia Venturini, and Matthias Eder
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MicroRNAs in Vascular Neointimal Lesion Formation .................... Chunxiang Zhang
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microRNA in Cutaneous Wound Healing ........................................... Chandan K. Sen and Sashwati Roy
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CpG Island Hypermethylation, miRNAs, and Human Cancer .............................................................................. Amaia Lujambio and Manel Esteller
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Microarray Profiling of microRNA Changes in Cells That Express HIV-1 Proteins ............................................................... Man Lung Yeung and Kuan-Teh Jeang
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microRNA-Associated Therapies ......................................................... Anne Saumet, Guillaume Vetter, Nicolas Cougot, Manuella Bouttier, Florence Rage, Khalil Arar, and Charles-Henri Lecellier
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The Use of RNAi to Elucidate and Manipulate Secondary Metabolite Synthesis in Plants ............................................................. George J. Wagner and Antoaneta B. Kroumova
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Index ................................................................................................................
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Chapter 1
Structures of MicroRNA Precursors Piotr Kozlowski**, Julia Starega-Roslan**, Marta Legacz, Marcin Magnus, and Wlodzimierz J. Krzyzosiak*
Abstract MicroRNAs are single-stranded regulatory RNAs of 18–25 nucleotide length generated from endogenous transcripts that form local hairpin structures. The processing of microRNA transcripts involves the activities of two RNase III enzymes Drosha and Dicer. In this study we analyzed structural features of human microRNA precursors that make these transcripts Drosha and Dicer substrates. The structures of minimal functional primary precursors (pri-microRNAs) and secondary precursors (pre-microRNAs) were predicted. The frequency, nucleotide sequence content and the localization of various structure destabilizing motifs was analyzed. We identified numerous pri-microRNAs which structures strongly depart from the consensus structure and their processing is hard to explain by the existing model of the Microprocessor complex. We also found a biased distribution of symmetric and asymmetric motifs along the pre-microRNA hairpin stem and an overrepresentation of bulges on its 5′ arm (p < 0.000001), which may have considerable functional implications. Keywords miRNA biogenesis, Dicer, Drosha, RNA structure prediction, pri-miRNAs, pre-miRNA structural motifs
1.1
Introduction
MicroRNAs (miRNAs) are a family of short single-stranded noncoding RNAs identified in many eukaryotes from simple organisms to humans [1, 8]. It is anticipated that hundreds of miRNAs regulate the expression of thousands of human genes [20]. MiRNAs regulate gene expression at the posttranscriptional
Laboratory of Cancer Genetics, Institute of Bioorganic Chemistry, Polish Academy of Science, Noskowskiego 12/14, 61-704 Poznan, Poland * Corresponding author: E-mail:
[email protected] ** These authors contributed equally to this work.
S.-Y. Ying (ed.) Current Perspectives in microRNAs (miRNA), © Springer Science + Business Media B.V. 2008
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level by programming the RNA induced silencing complex (miRISC) which interacts with the complementary sequences of mRNAs causing their translational inhibition or cleavage [25, 33]. Specific miRNAs were shown to be engaged in the variety of processes such as development, cell proliferation, differentiation and apoptosis [12]. The detailed cellular function of the majority of miRNAs still remains unknown. The primary transcripts of miRNA genes (pri-miRNAs) are generated by either RNA polymerase II [19] or RNA polymerase III [4]. The pri-miRNAs, which harbor a long stem and loop structure, are processed in the nucleus to shorter, approximately 60-nt hairpin precursors (pre-miRNAs) (Fig. 1.1A). The nuclear processing enzyme is ribonuclease Drosha [18] which acts together with the DGCR8 protein [17] within the Microprocessor complex [5, 7]. Drosha which is the RNaseIII enzyme usually leaves a 2 nt overhang at the 3′-end of pre-miRNA and defines one end of mature miRNA [2]. The pre-miRNAs are then exported to cytoplasm by Exportin-5 [22] and further processed to miRNA duplexes by another RNaseIII enzyme Dicer [3, 26, 34] which defines the other miRNA end. Thus, the two RNA processing steps reduce the stem of the primary precursor hairpin into its internal portion, which usually is the imperfect duplex containing a functional miRNA strand (Fig. 1.1B). The presence of structure imperfection within this duplex facilitates its non-miRNA strand to be later disposed from the miRISC using the “bypass” rather than the cleavage mechanism [6]. It is clear that the structures of miRNA precursors are instrumental for their proper recognition and correct cleavages by the processing complexes containing Drosha and Dicer. Therefore, to analyze the structural aspects of both the nuclear and cytoplasmic steps of miRNA biogenesis the structures of the primary and secondary miRNA precursors need to be established. This can be done either by experiment or computational structure prediction. The latter approach gives the reliable structures of miRNA precursors that were confirmed in most of the investigated cases by experimental analysis [13]. The computational approach is also much faster thus better suited for structure analysis on a large scale. In order to predict the secondary structures of miRNA precursors their nucleotide sequences need to be known first. These sequences, however, are not available in the existing miRNA databases. For the purpose of this study the sequences of pre-miRNAs were reconstructed from the sequences of mature miRNAs as described earlier [14]. To analyze the pri-miRNA sequences the concept of a “minimal” functional precursor was adapted [9]. The pri-miRNA precursors may be very long and as such they are not amenable for a detailed structure analysis. Therefore, we analyzed minimal pri-miRNAs which were considered to be the shortest fragments of primary precursors that contain all sequence and structure elements required to be functional substrates for a Microprocessor. The difference between our approach and that described earlier [9] was that not a single length but several different lengths of sequences harboring miRNA were analyzed. Both the reconstructed sequences of pre-miRNAs and arbitrarily selected sequences of minimal pri-miRNAs were then subjected to structure prediction and a detailed analysis of their secondary structures. We intended to learn more about the occurrence and localization of different types of secondary structure
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Fig. 1.1 Biogenesis of miRNA. (A) Steps of miRNA biogenesis. Proteins involved in substrate recognition and precursor processing are shown. (B) Schemes of the Microprocessor complex and RNase Dicer interacting with their substrates, pri-miRNA and pre-miRNA, respectively. Arrows indicate Drosha and Dicer cleavage sites. RIIIa and RIIIb indicate the RNase domains responsible for cleavage. dsRBD denotes the double-stranded RNA binding domain. The fragment which corresponds to miRNA is marked in gray. (C) Scheme of minimal precursor pri-miRNA. The SD (single-stranded–double-stranded) and SL (stem-loop) junctions, internal distances and analyzed region are indicated. SD is postulated to be the DGCR8 protein binding site
motifs within the precursor hairpin. A comprehensive inventory of such motifs was generated and analyzed in relevance to the Drosha and Dicer steps of miRNA biogenesis.
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1.2 1.2.1
P. Kozlowski et al.
Materials and Methods Analysis of Pri-miRNA Structures
The sequences of minimal pri-miRNAs of the four length variants 110, 130, 150 and 300 nt were precut from longer sequences withdrawn from the GenBank for all 461 miRNAs deposited in the miRBase database version 8.2. [8]. To obtain the minimal pri-miRNA sequence of selected length, the natural sequence extensions of the required and equal length were added to each end of the pre-miRNA. These 1,844 sequences were subjected to secondary structure prediction by free energy minimization using the Mfold program [36]. The suboptimality parameter was set at 5% which means that all structures that have the free energy of the formation (∆G) 5% higher than the lowest energy structure were also shown. For further analysis only the lowest energy structures were taken. The critical parameter analyzed in the predicted structures was the length of the base-paired region in which only minor structure distorting motifs were allowed to exist. Only those minimal pri-miRNAs were taken for further analysis which had the same structure of the analyzed fragment in at least the 150 and 300 nt length variants. There were 246 such precursors, and among them were 180 pri-miRNAs which had the same structure of the fragment of interest in all four length variants.
1.2.2
Analysis of Pre-miRNA Structures
Prior to the structure analysis of pre-miRNAs their nucleotide sequences were reconstructed following the rules described earlier [14]. Briefly, a one arm terminus of the precursor hairpin was defined by one end of the miRNA sequence and the second arm terminus was defined either by the miRNA* end or assuming the existence of the 2 nt 3′ overhang at the Drosha cleavage site. The secondary structures of the pre-miRNAs were predicted using Mfold as described above for pri-miRNAs. All secondary structure motifs present in the lowest free energy structures were catalogued in the format that included the number and sequence of nucleotides present in the specific motif, the motif orientation and its localization within the precursor structure. These motifs were classified into two major groups: symmetric internal loops (SL) and asymmetric internal loops (AL). The first group includes both single nucleotide mismatches SL1:1 and longer symmetric loops SL2:2, SL3:3 etc. The second group includes bulges of different length AL0:1, AL0:2, AL0:3 etc. and asymmetric internal loops ALX:Y where both X and Y are different from 0. Thus, each motif is denoted by two numbers separated by the colon. The number or sequence before and after the colon denotes nucleotides from the precursor 5′ precursor arm and 3′ arm, respectively. For example, the single nucleotide bulge “a” located in the 5′ arm is denoted either AL1:0 or a:0. The localization of the motif was numbered counting from the terminal nucleotide at the 5′ arm of the
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pre-miRNA. To unify the position numbering system for all types of structural motifs the position of nucleotide directly preceding any specific motif was assigned as the motif position.
1.2.3
Statistical Methods
For asymmetric motifs their equal distribution between two pre-miRNA arms was assumed (null hypothesis). To assess the potential deviation from this distribution the chi2-squared test was applied using Statistica (StatSoft, Tulsa, OK) or Prism v. 4.0 (GraphPad Software, San Diego, CA). To compare the distribution of symmetric vs. asymmetric loops in pre-miRNAs having a moderate and high number of stem structure distorting motifs the Fisher exact test was used for the 2 × 2 contingency table analysis (programs as above). Where applicable, the Bonferroni adjustment for multiple comparisons was used.
1.3 1.3.1
Results Pri-miRNA Structure and Drosha Step of miRNA Biogenesis
In agreement with the recently proposed model of the Microprocessor structure the DGCR8 protein binds to the base of the pri-miRNA hairpin stem and forms a platform for Drosha binding and precursor cleavage [9] (Fig. 1.1B). DGCR8 anchors to the single strand-double strand (SD) junction in the structure of pri-miRNA. The consensus structure of minimal pri-miRNA was established based on the analysis of the predicted structures of numerous human primary precursors and the structure prediction was performed using the 110 nt long sequence for each pri-miRNA [9]. In light of the fact that human pre-miRNAs span the length range of 42–82 nt [14], the length of 110 nt seemed insufficient for the reliable minimal pri-miRNA structure prediction. To minimize the risk of taking an incorrect structure into consideration besides the 110 nt also three longer sequences harboring miRNAs were used for structure prediction in our study. Such an approach was undertaken because the structures generated by computer programs used for RNA structure prediction by free energy minimization are more trusted if the same critical domains are predicted from the sequences of different lengths. The detailed analysis of the predicted structures of minimal pri-miRNA precursors was focused on the fragment localized between the pre-miRNA ends and SD junction (Fig. 1.1C), which was proposed to play a critical role in pri-miRNA recognition by the Microprocessor [9]. In the consensus minimal pri-miRNA structure this region spanned 11 bp, which equals to one helical turn of A-RNA. We wanted to find out whether the consensus minimal
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Fig. 1.2 Pri-miRNA stem length distribution. The length distribution of base-paired stems having either full base complementarity or only minor disruptions (small internal loops, bulges) within the analyzed fragment in 243 minimal pri-miRNA precursors. Above the graph schematic structures representing three classes of such precursors are shown. They differ in the length of the analyzed fragment
pri-miRNA structure is correct and find the most deviant structures which still remain substrates for the Microprocessor. It turned out from this analysis that there are indeed pri-miRNAs which secondary structures ideally fit to the consensus structure e.g. pri-miR-33 but there are also precursors which have the analyzed region either much shorter e.g. pri-miR-656 or much longer e.g. pri-miR-607. However, in the majority of the analyzed structures (62.6%) the SD junction was located 9–13 nt below the Drosha cleavage site which is in rough agreement with the consensus structure proposed by Han et al. [9] and confirmed by Saetrom et al. [28]. This region was shorter in 13% of the analyzed precursors and longer in 19.5% of the precursors (Fig. 1.2). It is difficult to fit such precursors, especially their extreme examples, into the presently accepted model of pri-miRNA processing by the Microprocessor complex (Fig. 1.1C). This may suggest that either some alternative models of Microprocessor architecture need to be considered or precursors having structures most deviant from the consensus are processed in an entirely different way.
1.3.2
Pre-miRNA Structure and Dicer Step of miRNA Biogenesis
The structural insights into Dicer function came from both biochemical studies [35] and crystallography [23, 24]. In Fig. 1.1B the commonly accepted model of a Dicer single processing center is shown [35]. Human Dicer is composed of several
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functional domains: the PAZ domain, which is used for high-affinity binding to the 3′ overhanging nucleotides of pre-microRNA, the helicase domain, the DUF283 subunit, the dsRNA binding domain and two catalytic RNase III domains that form the intramolecular dimer during pre-miRNA cleavage [24, 35]. Thus, Dicer functions as a molecular ruler and cleaves the pre-miRNA hairpin about two helical turns away from the hairpin base to produce duplexes containing 18–24 nt long miRNAs. It is intuitively understood that the length diversity of miRNAs has its source in the structural features of pre-miRNA hairpins which rarely contain perfectly base paired stems. Usually the single nucleotide mismatches, the larger symmetric internal loops, bulges and the asymmetric internal loops break the regularity of the pre-miRNA double helical stem structure and may influence both the efficiency of Dicer binding and specificity of precursor cleavage. Therefore, a detailed analysis of the predicted secondary structures of pre-miRNAs was performed to search for such structure distortions. Out of the 461 nucleotide sequences of human pre-miRNAs which were subjected to structure prediction nearly all (456) formed hairpins as the lowest free energy structures. In these precursor hairpins as many as 1,243 secondary structure motifs destabilizing stem structures were found altogether and the occurrence of various types of motifs in each arm of the hairpin stem is shown in Fig. 1.3. These motifs include 631 symmetric internal loops of various sizes including single nucleotide mismatches (SL) and a similar number (612) of asymmetric internal loops including bulges (AL) (Fig. 1.3B). This means that 2.73 motifs (0.97 mismatches,
Fig. 1.3 The frequency of structural motifs in stems of 456 analyzed pre-miRNAs. (A) The chessboard-like table shows in numbers the occurrences of each type of structural motifs identified in the predicted structures of miRNAs. These motifs are shown as pairwise combinations of unpaired nucleotides present in precursor 5′ and 3′ arm. E.g. symmetric loops SL are located diagonally and bulges along the 0 column. In this and the subsequent figures the motifs under consideration are shadowed. (B) The total number of symmetric loops (SL) including mismatches (SL1:1) and asymmetric loops (AL) including all types of bulges
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0.42 symmetric loops of different length (2–5), 0.96 bulges and 0.38 asymmetric internal loops) occur, on average, per analyzed pre-miRNA.
1.3.3
Symmetric Loops Including Single Nucleotide Mismatches in Pre-miRNA Structures
In cataloguing the structural motifs present in pre-miRNAs we have looked not only at their type and size but also at their nucleotide sequence and orientation within the precursor hairpin. This allowed us to count symmetric internal loops containing a different number of nucleotides and divide them into sequence and orientation- specific subgroups. Figure 1.4A shows the number of occurrences of single nucleotide mismatches and 2–5 nt long symmetric internal loops as well as the number of occurrences of different nucleotides and sequences present in these motifs. It is apparent that the frequency of symmetric loops decreases with their size. The single nucleotide mismatches are most frequent and account for almost 2/3 of the total number of symmetric internal loops. The largest is the 5 nt long loop identified only once in hsa-mir-196a-1. As shown in Fig. 1.4A all ten possible base combinations of single nucleotide mismatches are represented in the pre-miRNAs and 61 different combinations of 2 nt long internal loops. For the 3–5 nt long internal loops the number of different sequence classes is almost equal to the total number of such loops. This means that almost every symmetric internal loop formed by more than two adjacent nucleotides has a different sequence and there is no preference of any specific sequence within such loops. For the single nucleotide mismatches and 2 nt long internal loops we analyzed their distribution between different sequence classes (Figs. 1.4B, C, respectively). It turned out that the a:c and c:u are most frequent among the former and the least frequent is the c:c mismatch (Fig. 1.4B). The orientation analysis of single nucleotide mismatches shows that most of them are rather equally distributed in both orientations with the exception of the c:u in which u is more frequent on the 5′ arm and c on the 3′ arm (36 and 58 c:u and u:c mismatches, respectively). However, this distribution is only marginally significant (ch2; p-val = 0.02) and not significant after Bonferroni correction. The 2 nt internal loops are almost randomly distributed among sequence subclasses and clear ug:ug overrepresentation is only observed (19 occurrences) (Fig. 1.4C).
1.3.4
Asymmetric Internal Loops Including Bulges in Pre-miRNA Structures
As many as 437 bulges account for the majority of asymmetric loops identified in the analyzed pool of pre-miRNAs. These bulges vary in size from 1 nt (293 occurrences) to 11 nt (single occurrence) (Fig. 1.5A). The frequency of bulges decreases with bulge
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Fig. 1.4 The symmetric loops in pre-miRNAs. (A) The number of single nucleotide mismatches and symmetric loops containing different numbers of unpaired nucleotides within the loop. The number of different nucleotide combinations in symmetric loops (dark gray), and total number of symmetric loops (light gray). (B) The number of different types of mismatches with their orientation taken into account and cumulative number (inset). (C) As in (B) but for 2 nt symmetric loops
size with almost a perfect exponential correlation (r2 = 0.97). The distribution of the most frequent bulges (1–3 nt) between the precursor hairpin arms shows their overrepresentation in the 5′ arm (Fig. 1.5A). Testing the null hypothesis that bulges are equally distributed between the 5′ and 3′ arms we showed that the total overrepresentation of bulges in the 5′ arm is very significant (chi2 p-val < 0.000001). Individual chi2 p-values for the 1-, 2- and 3 nt bulges are 0.00002, 0.014 and 0.23 respectively. The nucleotides present in the single nucleotide bulges and sequences present in the
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Fig. 1.5 The bulges in pre-miRNAs. (A) (center) The total number of bulges containing a different number of nucleotides (1–11 nt). (right) For the most frequent 1–3 nt bulges the total number of bulges was split between two orientations 0:X and X:0 for bulges in the 3′ and 5′ arm of pre-miRNA, respectively. (B) The number of different nucleotides in single-nucleotide bulges. The 0:1 and 1:0 orientations are shown separately. (C) As in (B) but for different combinations of nucleotides in 2 nt bulges
2 nt bulges are shown in Fig. 1.5B, C, respectively. Among the former the most frequent is u and least frequent is g (Fig. 1.5B). The 2 nt bulges are almost equally distributed over sequence variants and almost all combinations of 2 nt sequences occur (except for gc) (Fig. 1.5C). The asymmetric internal loops containing a different number of unpaired nucleotides on each side constitute a smaller and more heterogenous group (177 occurrences). In this group the small motifs such as AL1:2 and AL2:1 are most frequent but single cases of large motifs e.g. AL6:4 and AL3:10 were also found. The analysis of their sequence contents did not reveal any significant preferences.
1 Structures of MicroRNA Precursors
1.3.5
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Localization of Structural Motifs Within the Pre-miRNA Hairpin Stem
The proper localization of structural motifs in pre-miRNAs may facilitate the adaptation of precursor structures to the interacting proteins of miRNA biogenesis machinery. It may create a suitable environment for interaction with the specific RNA binding motifs of proteins and serve as a code for structure-specific RNA recognition. Therefore we looked in this study also at the localization of structural motifs in the pre-miRNA hairpin stem. As shown in Fig. 1.6 there is no specific position in which the single nucleotide mismatches and symmetric internal loops would be either over represented or under represented (Fig. 1.6A, B). However, a clear trend is observed to decrease the frequency of these motifs in going from the precursor base towards the terminal loop. This trend is most clear for single nucleotide mismatches and 2 nt long symmetric internal loops. The number of longer symmetric loops (4 and 5 nt) is too low to see any trend (Fig. 1.6A). Interestingly, the opposite trend is observed for asymmetric motifs the frequency of which increases in the same direction (Fig. 1.6C).
1.4
Discussion
The bioinformatics survey of miRNA precursor structures which was performed in this study provides a comprehensive insight into the structural variety of both pri-miRNAs and pre-miRNAs. This insight may be considered as next step towards a better understanding of the role of RNA structure in miRNA biogenesis. The obtained gallery of predicted structures of miRNA precursors will guide the selection of specific precursors for a more detailed experimental analysis of their structures and studies of their interactions with Drosha and Dicer protein complexes. The structural features of the precursors of numerous known miRNAs will also help to refine algorithms used for the identification of novel miRNA genes in genomes. In addition, the structural information gathered in this study may be relevant to the process of RISC loading by miRNA/miRNA* duplexes that may retain the structure imperfections present within miRNA precursors. We catalogued the rich repertoire of secondary structure motifs destabilizing and distorting the stem structures of pre-miRNAs paying attention to the nucleotide sequences present within these motifs and motif localization. The detailed analysis of this data collection revealed that with some exceptions there are only minor preferences for specific sequences in the destabilizing motifs present in pre-miRNA structures. This means that protein complexes involved in miRNA biogenesis use structure rather than sequence code for precursor recognition. As shown in this study there are about 2.7 stem structure destabilizing motifs in the average pre-miRNA hairpin. The number of pre-miRNAs containing a different number of such motifs is almost normally distributed with extreme numbers being 0 and 7 motifs per premiRNA (Fig. 1.7A). Taking into account this distribution and assuming somehow
12 Fig. 1.6 The localization of structural motifs in the pre-miRNA hairpin structure. (A) The localization of mismatches SL1:1 and symmetric internal loops having a different number of nucleotides SL2:2 – SL5:5 within precursor structure. (B) The cumulative number of mismatched nucleotides along the pre-miRNA hairpin. (C) Localization of bulges within the precursor structure
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arbitrarily the ~5% threshold of pre-miRNAs with extreme numbers of structural motifs we divided all the analyzed pre-miRNAs into three classes: (1) having 0 structure destabilizing motifs (2%), (2) containing a moderate number (1–4) of such motifs (93%), and (3) harboring high number (5–7) motifs (5%). The analysis of the symmetric and asymmetric loops distribution revealed a gradual increase of asymmetric motifs with the total number of motifs in pre-miRNAs. When we compared the frequency of SL vs. AL in pre-miRNAs with a moderate and high number of motifs it showed significant excess of AL in pre-miRNAs containing a high total number of motifs (p-val = 0.0005) (Fig. 1.7B). This could reflect the compensatory effect to balance the structure distortion introduced by one bending motif by another. The biased distribution of bulges observed in this study consisted of their overrepresentation in the 5′arm of pre-miRNA. To validate this result we analyzed the distribution of bulges also in the group of “prototypical” pre-miRNAs recently distinguished by Tuschl’s group [16] on the basis of the precise miRNA 5′ end processing, sequence conservation and high number of putative target sites [16]. We have shown that in the “prototypical” group the overrepresentation of bulges in the 5′ arm is even higher than that revealed in the group of pre-miRNAs analyzed in our study (compare results shown in Fig. 1.8A with those in Fig. 1.5A). Although the number of “prototypical” miRNA precursors is smaller (266) than the total number of pre-miRNAs analyzed by us (456) the statistical significance of the disproportional distribution of bulges is even higher for the “prototypical” group. Contrary to that group the observed bias completely disappears in the group of
Fig. 1.7 Statistics of pre-miRNAs containing a high number, moderate number and no structure destabilizing motifs. (A) The number and frequency (inset) of pre-miRNAs having a different number of structure distortions in the hairpin stem. Assuming the ~5% threshold we distinguished three groups of miRNA precursors with a high number (5–7), moderate number (1–4) and no (0) stem distortion. (B) Distribution of SL and AL motifs in precursors containing a different number of structural motifs and frequency of SL and AL motifs in precursors having a moderate and high number of structural motifs (inset)
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“repeat-derived” miRNAs. Also the distribution of internal asymmetric loops observed in this study is in line with a lower tolerance of excessive nucleotides in the 3′ arm of pre-miRNA. Our analysis shows an overrepresentation of loops having a higher number nucleotides in the 5′ arm (Fig. 1.3A). However, the number of asymmetric internal loops is relatively small and this effect is not statistically significant. To find out whether the bulges overrepresented in the 5′ arm of pre-miRNA are equally distributed along the hairpin structure we compared the localization of the 5′ and 3′ arm bulges (Fig. 1.8B). It appears from this comparison that bulges in the 5′arm are not equally distributed but they tend to be clustered at two sites with maxima at nucleotide positions 11 and 18. These sites could be involved in the bending of the pre-miRNA structures and/or in interactions with specific protein domains. Comprehensive information on the distribution of various structural motifs in miRNA precursors will be also useful to fine-tune the algorithms used for the ab initio prediction of miRNA genes. Numerous algorithms have been developed to distinguish miRNA precursors from other hairpin structures encoded by genomes [10, 21, 30, 31]. These algorithms use different conservation, thermodynamic, sequence and structure parameters. The latter include some general parameters such as the length of the longest fully base-paired stem, terminal loop size, the number of nucleotides in the symmetric and asymmetric loops including bulges [15, 27, 29] as well as more specific structural characteristics such as frequency of triplet structure elements [11, 27, 32]. The results of our study show that there are also other highly
Fig. 1.8 The overrepresentation of bulges in the 5′-arm of prototypical pre-miRNAs. (A) As in (Fig. 1.5A) but separately for prototypical and repeat-derived classes of miRNA [16]. (B) Localization of bulges in the 5′-arm (black) or 3′-arm (gray) of pre-miRNA structure
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significant features of pre-miRNA structure that might facilitate miRNA gene prediction. These parameters include: strong overrepresentation of bulges in the 5′ arm of pre-miRNA, the opposite polarity of symmetric and asymmetric motifs distribution along the hairpin stem and increased contribution of asymmetric motifs when the total number of stem structure destabilizing motifs in pre-miRNAs increases. Acknowledgement This work was supported by funding under the Sixth Research Framework Programme of the European Union, Project RIGHT (LSHB-CT-2004-005276) and by the Ministry of Science and Higher Education, Grant No. N301 112 32/3910.
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Chapter 2
Small RNA Technologies: siRNA, miRNA, antagomiR, Target Mimicry, miRNA Sponge and miRNA Profiling Guiliang Tang1*, Yu Xiang2, Zhensheng Kang3, Venugopal Mendu1, Xiaohu Tang1, Xiaoyun Jia1, Qi-Jun Chen1, and Xiaoqing Tang4
Abstract The breakthrough discovery of RNA interference (RNAi) by Fire and Mello in 1998 has ushered in a new wave of RNA-based technological advances in the life sciences. Small RNAs, namely small interfering RNA (siRNA) and microRNA (miRNA), not only play key roles in down regulating gene expression, controlling growth and development, stress response, and various diseases, but also serve as essential tools for the study of gene functions. In this chapter, we provide an overview of the technological aspects of siRNAs and miRNAs and common methods for studying their functions. Keywords siRNA, miRNA, antagomiR, target mimicry, miRNA sponge, miRNA profiling
2.1
Introduction
RNA interference (RNAi) was the 2006 Nobel Prize winning discovery [135], although related research is still at an early stage and continuing at a rapid pace. RNAi technology has become one of the most important technological tools and is
1
Gene Suppression Laboratory, Department of Plant and Soil Sciences and KTRDC, University of Kentucky, Lexington, KY 40546, USA
2 Pacific Agri-Food Research Centre, Agriculture and Agri-Food Canada, Box 5000, 4200 Highway 97, Summerland, BC V0H1Z0, Canada 3 College of Plant Protection, Shaanxi Provincial Key Lab of Molecular Biology for Agriculture, Northwest Agriculture and Forestry (A & F) University, Yangling, 712100, China 4 Department of Molecular and Cellular Biochemistry, University of Kentucky College of Medicine, 741 South Limestone, Lexington, KY 40536, USA
* Corresponding author: Phone: 856 257 1594; Fax: 859 323 1077; E-mail:
[email protected] S.-Y. Ying (ed.) Current Perspectives in microRNAs (miRNA), © Springer Science + Business Media B.V. 2008
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now widely applied in almost every research aspect of modern biology. The key elements in RNAi are the small RNAs of ∼21 nucleotides (nt), termed small interfering RNAs (siRNAs) [22–24, 39, 138]. SiRNAs regulate their target genes by binding in a sequence-specific manner to target gene transcripts (e.g. mRNAs), and inducing degradation of target RNAs or blocking their translations [20, 138]. The simple structure of siRNAs allows them to be easily generated in large quantities by chemical synthesis [22]. Synthetic siRNAs, as powerful reagents, bypass their biogenesis steps in RNAi and induce potent and specific silencing of any gene of interest in cells [22]. MicroRNA (miRNA), the siRNA cousin, is produced by RNAi-like mechanisms or miRNA pathways [3], and plays a key role in gene regulation, cell developmental control and various disease development [7]. More than 450 miRNAs encoded by the human genome have been identified and are predicated to regulate the expression of about one third of human coding genes [71, 130]. The development of many types of cancers or diseases is related to the abnormal expression or loss of certain miRNAs [11, 12, 17]. The global miRNA expression patterns of specific organs, tissues or cells can serve as an miRNA atlas or as biomarkers for the study of miRNA functions [65]. As a result, one can potentially reverse metastatic cancers by blocking the abnormally-expressed miRNAs or by reintroducing the lost miRNAs during cancer progression [80]. Much evidence indicates that many miRNAs function not only individually but also coordinately in gene regulation or in specific disease development [41]. Thus, this creates a new direction for studying a specific disease at the cellular level by simultaneously manipulating multiple miRNAs. Based on the study of the structures of miRNAs, artificial miRNAs have been developed as powerful tools for potently silencing genes in plants and animals [1, 89, 94, 101, 139, 140]. Compared to previous RNAi technologies, artificial miRNAs more accurately and specifically silence genes of interest and reduce off-target effects. More than 5,000 miRNAs have been identified from various organisms (http://microrna.sanger.ac.uk/) [32, 33], but very few of them have been functionally analyzed. It remains a big challenge to discover the functions of most miRNAs currently stored in the database. The emergence of new technologies, such as highthroughput miRNA array, antagomiR [59], miRNA target mimicry and miRNA sponge [18, 21, 30] provide new tools to understand how, where and when miRNAs are generated and function in specific tissues, cells and organisms. This chapter gives an overview of these different small RNA technologies and their applications.
2.2
The Basic Biology and Chemistry of siRNAs and miRNAs and Their Related Working Mechanisms
SiRNA and miRNA are ~21–23 nt small RNAs produced in cells via a series of enzymatic steps. Long double-stranded RNA (dsRNA) or stem-loop structured RNAs introduced or transcribed in cells are first processed into siRNA or miRNA duplexes by a dsRNA specific RNase-III family of enzymes termed Dicers [137].
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The small RNA duplexes are characterized by 2-nt overhangs at the 3′ ends with specific chemical end structures of a monophosphate at the 5′ end and a hydroxyl group at the 3′ end [44, 115]. In addition to differences in biogenesis, the siRNA duplex is a perfect duplex of Watson-Crick base pairs while the miRNA duplex is often an imperfect duplex with mismatches or bulges [114]. Both RNA duplexes need unwinding to form effector complexes: RNA-induced silencing complexes (RISCs) for their functionality. The key protein component of the effector complex is the Argonaute (AGO) protein of an evolutionally conserved protein super family [14]. The end structures of the small RNA duplex play a pivotal role in the assembly of the effectors or RISCs [76, 108]. After RISC assembly and maturation, specific gene transcripts, or messenger RNAs (mRNAs), are recognized and bound by the small RNAs on the RISCs based on sequence complementarities, leading to site-specific cleavage or translational repression of the mRNAs [28]. The elucidation of siRNA or miRNA chemical structures provides chemical approaches to generate them in large quantities for RNAi applications. Although single-stranded siRNA can be assembled into RISCs [81], synthetic siRNAs or miRNAs usually need to be double-stranded to be recognized by Dicer for initiating effective RISC assembly [77]. Thus, two short complementary RNA strands need to be synthesized and annealed to form siRNA or miRNA duplex characteristics with bona fide siRNA or miRNA end structures. In Drosophila, structurally distinct siRNA and miRNA duplexes are recognized by different Dicer proteins (DCR-1 and DCR-2) and sorted into distinct AGO proteins (AGO-1 and AGO-2) [29, 69, 91, 117]. DCR-2 sorts the siRNA duplex to be assembled into AGO-2 while DCR-1 sorts the miRNA duplex into the AGO-1 protein complex. Sometimes the two sorting systems are interchangeable based on siRNA or miRNA duplex structures or nucleotide compositions [29, 117]. SiRNA or miRNA duplexes can be divided into two types according to their end structures and thermodynamic stability: symmetric and asymmetric siRNA or miRNA [114]. A symmetric siRNA or miRNA assembles into two kinds of RISCs with either a sense strand or an antisense strand and can potentially interact with two different complementary target mRNAs for regulation [114]. However, siRNAs or artificial miRNAs are normally designed to target one specific gene transcript rather than two. Thus, symmetric siRNAs or miRNAs have a higher chance of targeting unwanted mRNAs due to their ability to target two different gene transcripts, leading to off-target effects. In contrast, asymmetric siRNAs or miRNAs preferentially favor only one specific strand of the small RNA duplex assembled into a RISC while the other strand is excluded from RISCs and is subsequently degraded [102]. This allows a maximum assembly of one specific strand of small RNA duplex into RISC components, considerably reducing off-target effects coming from the RISC assembled by the unwanted strand. Intriguingly, a major fraction of endogenous miRNAs are structurally asymmetric and display high specificity in the regulation of their target genes [50, 102], although a substantial number of miRNAs seem to have dual functions coming from each strand of the miRNA duplex (personal communication with Eric Lai). The asymmetric character seems a result of natural selection to reduce unwanted
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off-target effects for fine control of development during evolution. The asymmetric structure of endogenous miRNAs has been successfully adopted to design highly efficient artificial miRNAs to target genes of interest in plants and animals [1, 89, 94, 101, 139, 140]. Specific programs have also been developed to help design highly specific artificial miRNAs [101]. How can asymmetric siRNAs or miRNAs ensure that one specific strand is going into the RISC and excluding the other strand to prevent off-target effects? Recent research results indicate that RISC assembly and activation involve strand selection and exclusion by the RISCs. In the case of siRNAs, guide strand-associated RISCs exclude the passenger strand by guide-associated RISC directed cleavage of the passenger [70, 82, 95]. This cleavage has the same characteristics of RISCdirected target mRNA cleavage. That is, the cleavage site on the passenger strand is between the bases 10 and 11 from the 5′ end of the guider siRNA strand [24]. However, this cleavage depends on several prerequisites [82]. First, the AGO protein on the RISC must have “slicer” (endonuclease) activity. Second, there must be no mismatches or bulges between the guide strand and the passenger strand around the cleavage site. Thus, miRNA duplexes, which naturally have bulges or mismatches around the potential cleavage sites, likely employ other mechanisms to exclude and eliminate the passenger strand. This mechanism should be also true for RISCs on which the slicer activity is missing. In such cases, it remains unknown how the passenger strands are degraded after their exclusion from the RISCs, rather than cleaved by the slicer.
2.3
MiRNA Functional Analysis and miRNA Inhibitors
Like transcription factors, miRNA-directed regulation of post-transcriptional gene expression is wide-ranging [41]. First, the expression of many transcription factors themselves are regulated by miRNAs in plants and animals. Second, miRNAdirected gene regulation seems more extensive in animals than in plants due to distinct working mechanisms. One third of the human protein-coding genes are predicted to be regulated by different miRNAs [71, 130], but relatively very few miRNA-target interactions have been experimentally validated. Most identified or predicted miRNAs are functionally unknown. Thus, the study of miRNA functions constitutes a unique aspect of miRNA genomics. Due to the small size of miRNAs and their functions associated with specific target genes, functional analysis of miRNAs is substantially different from coding genes and as a result, various methods have been developed. An earlier approach to the study of miRNA targets was focused on computational predications based on the base-pairing conditions between miRNAs and their target genes [16, 51, 56, 96, 100, 106, 123, 124, 131, 134, 141]. The completion of the whole genome sequencing of various organisms expedited the discovery of new miRNA genes and their targets by a computational approach. The regulatory roles of miRNAs are reflected in the cellular functions of miRNA target genes.
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The prediction that one-third of human protein-coding genes are miRNA targets was mainly based on the assumption that a short stretch of “seed” region (~7 base pairs of the position 2 to 8 from the 5′ end of the miRNAs) between miRNAs and the 3′ untranslated region (UTR) of their targets is sufficient for regulation [71, 72, 130]. In contrast, regulation of target gene transcripts in plants by miRNAs seems predominantly reliant on extensive sequence complementarities between the miRNAs and their target genes. Thus, the number of predicted miRNA targets in plants is much more limited [98]. The outcomes of the interactions between plant miRNAs and their targets, in most cases, are the cleavage and degradation of the target mRNAs. Thus, validation of plant miRNA targets is relatively simple and can be done by assaying for target cleavage in vitro and in vivo. The first validations of miRNA-target interactions were reported in plants [78, 115]. Two experimental approaches were established for these validations: direct visualization of the target cleavage in vitro and cloning of the target cleavage products in vivo by 3′ or 5′ RACE-PCR. Traditional wheat germ extracts or newly established maize germ extracts (G. Tang, 2007) are convenient cell-free systems for miRNA target validation. Both systems contain abundant endogenous miRNAs that have already been loaded on the RISCs, and exogenous target mRNAs are readily cleaved by the existing miRNA-associated RISCs. However, these cell-free systems have limitations when validating miRNA-target interactions whose corresponding miRNAs do not exist in wheat or maize germ extracts. In vitro programming of active RISCs by synthetic miRNA duplexes in plant systems needs further exploration [83, 93, 115]. In contrast, Drosophila embryo extracts are capable of RISC assembly programmed by various kinds of small RNAs and can thus serve as a platform for animal miRNA target validations, as well as a useful heterologous system for validation of plant miRNA targets [38]. The validation of animal miRNA-target interactions is not trivial. First, most miRNA targets in animals are not directly cleaved but rather translationally repressed by miRNA-associated RISCs. Therefore, the validation needs to be conducted at the protein level. Detection of the change in protein expression of the target genes using specific antibodies is preferred for such validation [143]. Alternatively, reporter genes are often fused with the 3′ UTR of the target mRNAs for the purpose of validating translational repression by miRNAs [58]. A mammalian cell-free system was recently developed to recapitulate let-7 miRNA-directed translational repression in vitro, which will be useful in the validation of animal miRNA-target interactions [121]. This in vitro system was established with extracts from HEK293F cells transfected with expression vectors that contain genes encoding various miRNA pathway components, such as Dicer, TRBP2, Argonaute2 and GW182. This system is capable of processing chemically synthesized let-7 miRNA precursors into mature let-7 that is likely to be further assembled into RISC to direct translational repression. Based on this system, Wakiyama et al. found that let-7 miRNP complexes induced the deadenylation of the let-7 target mRNAs and the abolishment of cap-poly(A) synergy, leading to target mRNA translation blockage [121]. These in vivo and in vitro approaches are often used together to validate animal miRNA targets.
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Approaches to inducing a loss of function of miRNAs represent a powerful functional genomics tool. Traditional screening for mutation of miRNA genes has not proved very successful. This is because miRNAs are often encoded by multigene family members and the loss of function of one miRNA member is often obscured by redundant functions of other miRNA members that have almost identical sequences and the ability to bind and regulate the same target gene transcripts. Fortunately, different methods and strategies have been developed to block the regulatory functions of all members of any miRNA family. In vitro chemically modified miRNA inhibitors, such as ‘antagomirs’, or in vivo target mimicry in planta and miRNA sponge in mammalian cells have recently proved to be effective in blocking functions of specific miRNA families [21, 30, 59]. Chemically modified oligonucleotides have been widely used in the study of the loss-of-function of miRNAs. Based on antisense strategy, oligonucleotides complementary to the miRNAs act as competitive inhibitors of endogenous target mRNA binding to the miRNAs, leading to a suppression of miRNA functions. These have been developed and demonstrated to be very specific and potent inhibitors of targeted miRNAs [45, 59]. The major modification in antagomirs is 2′-O-methylation of the ribose, sometimes combined with other kinds of modifications such as phosphorothioate linkage near 5′ and 3′ ends and a cholesterol-moiety conjugated at 3′ end. Antagomir, usually 21–33 nt in length, sequence-specifically binds to specific target miRNAs through base-pairing [45, 59]. A traditional modified antisense oligo, such as morpholino that was previously used to knock down the expression of protein coding gene transcripts, also successfully knocked down specific miRNAs by binding to the miRNAs or their precursors [54]. These chemically modified miRNA antisense RNAs can effectively compete with miRNA target mRNAs by a stronger bind to their specific target miRNAs on the miRISCs, resulting in inhibition of the miRNA activities. Antagomirs also induced the degradation of the targeted miRNAs with as of yet unknown mechanisms [57, 59]. Antagomirs were delivered to mouse tissues via intravenous injections, absorbed by tissues, and were highly resistant to various RNases in cells. It was shown that antagomirs specifically inactivated multiple target miRNAs in various mouse tissues for over 20 days following a single intravenous injection, resulting in changes in the abundance of distinct target mRNAs [57, 59]. The introduction of antagomirs against specific miRNAs in cells will release the repression of the bona fide miRNA target mRNAs from translation into proteins, thus indirectly validating the miRNA targets. The strategy of target mimicry to block miRNA functions was enlightened by a study of the interactive relationships between the phosphate (Pi) starvation-induced miR-399 and its naturally occurring target RNA transcripts from IPS1 gene in Arabidopsis thaliana [30]. IPS1 contains a 23-nucleotide motif that is almost complementary to miR-399 but with a mismatched loop at the expected miRNAdirected cleavage site. Over-expressed IPS1 RNAs can bind to mature miR-399 associated RISCs, and prevent miR-399 mediated cleavage of the target mRNAs, including PHO mRNA. Mutation of the IPS1 motif to be perfectly complementary to miR-399 abolished IPS1 inhibitory activity on miR-399, indicating that miR-399 associated RISCs are highly efficient and multiple-turnover enzymes to cleave their
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perfectly complementary targets, but may be stuck in the interaction between RISCs and their non-cleavable targets. This research not only revealed the phenomenon that miRNA activity in planta can be regulated by its mimic non-coding RNA, but also provided a genetic tool for the miRNA functional analysis. That is, introduction of an artificial non-cleavable miRNA target mimic is capable of knocking down complementary miRNA activity in vivo. Using this strategy and the IPS gene backbone [30], successfully inhibited the activities of the miR-156 and miR-319 separately by over-expressing target mimics with non-cleavable sites for these miRNAs, respectively, in Arabidopsis thaliana. The principle of ‘miRNA sponges’ developed by [21] is similar to target mimicry of miRNAs described in plants. Artificial target RNAs are designed to contain several tandem complementary binding sites to the miRNA of interest, but with a bulge or mismatch in the RISC cleavage site, and are genetically engineered to be stably expressed in mammalian cells. These artificial RNAs, like sponges, absorb a high level of their complementary miRNAs and release the translational repression of the bona fide targets by the miRNA-associated RISCs (miRISCs). The mismatch in the cleavage site prevents the decoy RNA from being degraded by miRISCs, but binds more firmly to the target miRNA loaded in RISC, sequestering it away from its bona fide mRNA targets in the cell. These miRNA sponges are experimentally proven to function as highly competitive miRNA inhibitors and depress miRNA targets effectively in mammalian cells. In addition, the sponges can be designed to bind effectively to multiple miRNAs that contain the same “seed’ region (position 2–8 from the 5′ end of the miRNA) [21].
2.4
SiRNA and miRNA Vectors and Their Application in Gene Silencing
Various siRNA vectors have been developed for knocking down genes in plants and animals since the discovery of RNAi [2, 26, 49, 68, 85, 97, 109, 110, 126, 128]. SiRNA vectors are able to generate siRNAs transiently or consistently from doublestranded RNAs or hairpin/stem-loop structure via Dicer enzymes targeting specific gene transcripts in various tissues or cells. However, not all siRNA vectors work well for both plants and animals. For example, the most popular siRNA vectors for gene silencing in plants are inverted repeat sequences coupled with a linker or a spliceable intron between the two repeats to form long (>100 bp) RNA hairpins [27, 62, 107, 125]. Yet, these vectors are not applicable in animal cells because the long doublestranded RNAs (dsRNA) generated from these vectors trigger cellular interferon pathways and lead to non-specific programmed cell death. Consequently, vectors that produce short hairpin RNAs (shRNAs) but which rarely trigger the interferon pathway were developed and have become widely used in animal studies [110, 129]. Similar shRNA vectors were also successfully applied toward directing gene silencing in plants [79]. Most shRNA vectors currently use an RNA polymerase III (Pol III) promoter, usually U6 or H1, and a pol III terminator (a stretch of thymidines)
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to transcribe a short hairpin structure with a stem of 19–29 bp and a short loop of 4–10 nt. The shRNA structures are further recognized and cleaved by Dicers to produce a large amount of siRNAs in cells to target specific gene transcripts. Later on, with the discovery of miRNA genes that produce highly specific short miRNAs, earlier vectors that produce short hairpin structured siRNAs were further modified by using miRNA backbones [9]. The new vectors adapted the flanking and loop regions of endogenous miRNAs to express miRNA-like primary transcripts (pri-miRNA-like RNAs) using pol-II promoters and terminators. These modified miRNA-like small RNA vectors have been demonstrated to be more effective in gene knock-down by many folds, indicating the miRNA backbones are structurally predisposed to produce more effective small RNAs for gene silencing [9]. The most significant discovery that helped to revolutionize small RNA vector technology was the discovery of the asymmetric structures of siRNAs [50, 102]. The realization that most endogenous miRNAs are structurally asymmetric immediately prompted the birth of second-generation RNAi vectors, artificial miRNA vectors [1, 89, 94, 101, 139, 140]. The major difference between miRNA backboned siRNA vectors and artificial miRNA vectors is found on the stem region of the stem-loop structured RNAs produced by both kinds of vectors. While the stem regions of the stem-looped structured RNAs produced by the miRNA backboned siRNA vectors are perfect Watson-Crick base pairs, the ones from the miRNA vectors are often designed to be imperfectly matched with mismatches, GU wobbles, and bulges. The biogenesis of artificial miRNAs produced by miRNA vectors strictly follows the miRNA pathways distinct from the RNAi pathways in cells. Thus, artificial miRNA vectors can be used not only to silence most protein coding genes, but also to knock down genes encoding the enzymes/proteins of RNAi pathways. Today, artificial miRNAs are widely used for gene silencing in both plants and animals. Compared to siRNA vectors, artificial miRNA vectors have significant advantages, including high specificity, fewer off-target effects, tissue-specific expression and almost no side effects. In contrast to the vectors that are used to silence individual genes, various modified vectors that direct simultaneous silencing of multiple genes have also been developed. For example, modified multi-hairpin structures of miR-30 have successfully knocked down multi-genes in a single construct [111]. We expect future gene silencing vectors to be more powerful in fine-tuning silencing of genes with subtle differences for various therapeutic purposes.
2.5
MiRNA Profiling and miRNA Biomarkers
Over 5,000 miRNAs from 44 organisms have been identified/predicted and stored in the miRBase registry (http://microrna.sanger.ac.uk/) [32, 33]. Most of them are not characterized by function. Determination of miRNA functions and miRNA-target interactions is therefore a long-term objective. Roles of miRNAs in post-transcriptional gene regulation are presumed to be extensive. In humans, an individual miRNA is predicted to regulate hundreds of coding gene transcripts [10, 71, 130]. Furthermore,
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transcripts from a single gene can be regulated by multiple miRNAs in a coordinated manner [25, 42, 48]. Analysis of miRNA expression profile is thus important in elucidating roles of specific miRNA or miRNA cohorts in the regulation of their target gene transcripts. Like the expression of mRNAs, the expression of miRNAs varies considerably from different cells, tissues, organs and species [63, 64]. The big challenge of studying miRNAs is how to effectively analyze thousands of miRNAs simultaneously for a limited amount of given samples. Traditional microarray technology has succeeded in analyzing entire protein-coding gene transcriptomes in various organisms. Similarly, various laboratories and companies have adopted the traditional mRNA array platform for the analysis of miRNAs over the last few years, allowing thousands of miRNAs to be analyzed from samples including plants and animals [6, 13, 15, 36, 47, 55, 67, 73–75, 84, 86, 87, 103–105, 113, 116, 122, 142]. The technologies to be used for miRNA array are not trivial. Compared to array analysis for mRNA expression, miRNA arrays usually involve much more complicated procedures due to the small size of the miRNAs and the lack of a conserved 3′ end for easy sample labeling. These complicated steps include the ligation of RNA adapters to the miRNAs, RT-PCR amplification and T7 RNA polymerase transcription [19, 87]. Current non-isotope miRNA array platforms involve these complicated techniques and require special skills; moreover, process-related systematic biases are unavoidable [19]. To simplify these steps, we have recently optimized the conditions for an earlier version of an isotope labeled miRNA array platform [55, 86], and further developed this system demonstrating its use in mouse miRNA analysis [116]. This optimized miRNA array platform is characterized by several unique features: (1) a careful selection of miRNAs for probe design to reduce potential cross-hybridization between different probes; (2) isolation of small RNAs of 15–28 nt using a 15% sequencing gel to avoid interference of signals between pre-miRNAs and pri-miRNAs; (3) direct labeling of the isolated small RNAs at their 5′end by introducing isotope-labeled phosphates to avoid using adaptors and biased PCR amplification, and directly hybridizing the labeled small RNAs to the membrane containing arrayed miRNA probes. Results can be output as a visual miRNA atlas reflecting the bona fide level of miRNAs in cells; and (4) introduction of a new way of data normalization by using Northern blot analysis of a constitutively expressed miRNA for initial data adjustment, and by a set of external controls for the evaluation of process-related loss of signal and quantification of endogenous miRNAs. Application of various miRNA array platforms reveals numerous miRNA biomarkers for a variety of human diseases including various kinds of cancers. The expression changes of these miRNA biomarkers indicate changes from normal to abnormal genetic and physiological conditions. For example, a specific spectrum of miRNAs including miR-23, -24, -26, -27, -103, -107, -181, -210, and -213 were induced in neoplastic cells under a hypoxic environment compared to normal conditions [61]. Similarly, comparisons of miRNA profiles of tumor and normal tissues have revealed distinct miRNA biomarkers for various tumor cells over the last a few years [11, 12, 46, 60, 112, 120, 132]. Exploration of these miRNA biomarkers will be particularly useful in early diagnostics of human diseases such as cancers,
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diabetes, and Alzheimer’s disease. Eventually, detailed miRNA atlases of humans, animals and plants will be available with the help of small RNA deep sequencing and miRNA array platforms for a wide variety of applications.
2.6
Conclusions and Perspectives
In conclusion, the study of RNAi and miRNAs has led to a number of small RNA technologies that are and will continue to be extremely useful in the study of small RNAs, individual gene functions, functional genomics, and various biological questions in both plants and animals. The dissection of RNAi, miRNA, and other small RNA pathways is only the beginning. For most organisms, the detailed maps or atlases of small RNAs have not been completed or even started. The small RNA networks in gene regulation are still obscure, and the entire picture of gene regulation by small RNAs and their detailed regulatory steps is a long-term goal. With a deeper understanding of the roles and mechanisms of small RNA-directed gene regulation, new technologies of using small RNAs will continue to be developed and established. We expect that such small RNA technologies will be expanded from the current siRNA or artificial miRNA directed post-transcriptional gene regulation to transcriptional gene regulation, for example, small RNA directed chromatin modifications or DNA methylation, to modulate the expression of any gene of interest. Small RNA technologies will not only used as tools to silence genes of various pathways but also genes of small RNA pathways themselves. For example, since RNAi and miRNA generally belong to different pathways that composed of different sets of enzymes, we expect that RNAi technology will be used to study functions of genes of the miRNA pathway, and vice versa. In addition to siRNAs and miRNAs, other kinds of small RNAs, such as trans-acting siRNAs (ta-siRNAs) [31, 40, 43, 90, 119, 127], repeat-associated small interfering RNAs (rasiRNAs) [37, 52, 53, 92, 118], and piwi-associated siRNAs (piRNAs) [4, 5, 8, 34, 35, 66, 88, 99, 133, 136], may also be able to be used for silencing of genes of interests. Acknowledgements G.T. is supported by the Kentucky Tobacco Research and Development Center (KTRDC), the USDA-NRI grants 2006-35301-17115 and 2006-35100-17433, and the NSF grant MCB-0718029, Subaward No. S-00000260.
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Chapter 3
RNA Interference Expression Vectors Based on miRNAs and RNA Splicing Akua N. Bonsra, Joshua Yonekubo, and Guangwei Du*
Abstract RNA interference (RNAi) has emerged as a powerful tool in basic research and therapeutics by silencing the expression of specific target genes. RNAi occurs naturally within cells to regulate gene expression at the post-transcriptional level. The development of reliable RNAi vectors encoding artificial and natural miRNAs would be useful tools for many RNAi applications. Here, we describe two new RNAi vectors, designated pSM155 and pSM30, that take into consideration of miRNA processing and RNA splicing by placing the miRNA-based artificial miRNA expression cassettes inside of synthetic introns. These vectors significantly improved the expression of a co-expressed enhanced green fluorescent protein (EGFP) marker and also provide a simplified cloning method. We discuss the advantages of these vectors, their potential applications, and concerns in using miRNA-based vectors. Keywords microRNA, mRNA, small-hairpin RNA, RNA interference, RNA splicing, intron
Abbreviations RNA interference, RNAi; small interfering RNAs, siRNAs; smallhairpin RNAs, shRNAs; microRNA, miRNA; primary miRNAs, pri-miRNAs; miRNA precursor, pre-miRNA; nucleotide, nt; enhanced green fluorescent protein, EGFP; oligonucleotide, oligo.
Department of Integrative Biology and Pharmacology, University of Texas Health Science Center at Houston, Houston, Texas 77030, U.S.A. * Corresponding author: E-mail:
[email protected] S.-Y. Ying (ed.) Current Perspectives in microRNAs (miRNA), © Springer Science + Business Media B.V. 2008
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Introduction
RNA interference (RNAi) has emerged as a powerful research tool for the silencing of specific target genes and shown great potentials in therapeutics [4]. RNAi occurs naturally within cells to regulate gene expression at the post-transcriptional level. Scientists are able to mimic and manipulate this process in mammalian cells by introducing small interfering RNAs (siRNAs) or by transfecting DNAbased vectors encoding short hairpin RNAs (shRNAs) [1, 10]. Recent strategies for gene silencing come from new RNAi vectors based on microRNAs (miRNAs) [2, 11, 12]. In eukaryotes, primary miRNAs (pri-miRNAs) are transcribed by RNA polymerase II in the nucleus, capped at the 5′ end and polyadenylated at the 3′ end [3, 7]. Pri-miRNA is processed by the nuclear microprocessor complex where an RNase III enzyme, Drosha, cleaves it at specific cites surrounding the hairpin [3, 7]. The resultant 70–90 nt hairpin has a distinct 2 nt 3′ overhang that is recognized by Exportin-5, which transports this precursor miRNA (premiRNA) out of the nucleus. A cytolasmic RNase III enzyme, Dicer, then acts on the pre-miRNA to produce a ∼22 nt double-stranded miRNA. The duplex separates into single-stranded mature miRNAs and enters the RNA-induced silencing complex (RISC). The miRNA within the complex guides it to the target mRNA, which it binds then, depending on complementarity, triggers for degradation or translation inhibition [3, 7]. Because of the growing need for understanding of the functions of miRNAs and the many beneficial uses of RNAi in scientific research and therapeutic applications, the development of reliable vectors encoding artificial and natural miRNAs has become critical. This chapter will describe the development of new miRNA/ shRNA expression vectors designed by our group which takes advantage of naturally occurring mechanisms, summarize protocols on using these vectors [6], and discuss their potential applications in basic research and therapeutics.
3.2 3.2.1
Materials and Methods General Reagents and Antibodies
Cell culture media, Dulbecco’s Modified Eagle Medium (DMEM), Opti-MEM-I, and LipofectAMINE Plus were from Invitrogen (Carlsbad, CA, USA). The GeneRuler 1 kb DNA Ladder Plus was from Fermentas (Glen Burnie, MD). All other reagents were of analytical grade unless otherwise specified. The rabbit polyclonal anti-PLD2 was kindly provided by Y. Banno (Gifu University of Tokyo, Gifu, Japan). Rabbit anti-green fluorescent protein (GFP) was from Abcam (Cambridge, MA, USA). Monoclonal anti-α-tubulin was from Sigma-Aldrich (St Louis, MO, USA). Goat anti-mouse and anti-rabbit IgG conjugated to Alexa
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680 were from Invitrogen. Goat anti-mouse and anti-human IgG conjugated to IRDye 800 were from Rockland Immunochemicals (Gilbertsville, PA, USA).
3.2.2
Construction of pSM155 and pSM30
The construction of pcDNA3.1-mCherry, pSM155, and pSM30 have been described [6]. Briefly, a synthetic exon and intron is placed between the cytomegalovirus (CMV) promoter and EGFP in the pEGFP-N1 vector (Clontech, Pal Alto, CA), to generate pEGFP-N1-Intron. The expression cassettes for miR155 or miR30 were then cloned into pEGFP-N1-Intron to get pSM155 and pSM30. Two inverted BsmBI sites were introduced into both vectors to facilitate subsequent insertion of artificial miRNAs. The oligos for candidate sequences also contained cohesive ends for a simplified cloning strategy. The plasmid maps were generated using VectorNTI from Invitrogen.
3.2.3
Synthesis of Oligos
3.2.3.1
Synthesis of Oligos for pSM155 (An Example Is Shown in Fig. 3.3A)
A. Generating the top oligo sequence. To generate the top oligo sequence, combine these elements (from 5′ end to 3′ end): start with 5′ TGCTG, reverse complement of the 21 nt sense target sequence (this is the mature miRNA sequence), add GTTTTGGCCACTGACTGAC (terminal loop), and add nucleotides 1–8 (5′-3′) of sense target sequence and nucleotides 11–21 (5′-3′) of the sense target sequence. B. To generate the bottom oligo sequence, perform the following steps: Remove 5′ TGCT from the top oligo sequence (new sequence starts with G), take the reverse complement of the sequence from step 1 and add CCTG to the 5′ end of the sequence from step 2.
3.2.3.2
Synthesis of Oligos for pSM30 (An Example Is Shown in Fig. 3.3B)
A. To generate the top oligo sequence, combine these elements (from 5′ end to 3′ end): start with 5′ AGCG, add a 22 nt sense target sequence, change the first nt to one which does not anneal to the last nt in the anti-sense, e.g. C to A, add TAGTGAAGCCACAGATGTA (terminal loop) and add nucleotides of the antisense target sequence. B. To generate the bottom oligo sequence, perform the following steps: remove 5′ AGCG from the top oligo sequence (new sequence starts with G), take the reverse complement of the sequence from step 1 (this is the mature miRNA sequence, and add GGCA to the 5′ end of the sequence from step 2.
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Cloning of Artificial miRNAs
1. Preparation of vectors: Cut 1–2 µg of pSM155 or pSM30 in 15 µl NEB (New England Biolabs) restriction Buffer 3 and BsmB I at 55 °C for 1 h. Run the digested vectors on 0.8% agarose gel and purify them using QIAEX II gel extraction kit following the manufacturer’s instructions (Qiagen, Valencia, CA). 2. Annealing of oligos (25 uM final concentration): Dissolve oligos with distilled water (100 uM stock concentration). Take 5 µl from each oligo (top and bottom) and add 8 µl water and 2 µl 10 × NEB restriction buffer (we usually use Buffer 3). Boil the oligos for 4 min. Leave the denatured oligos at room temperature for 15 min and then move them to 4 °C for 10–15 min. Dilute 2,500-fold to get 10 nM double-stranded oligos (250-fold dilution with water, quickly followed by 10-fold with 1 × NEB Buffer 3; or add 0.2 µl of annealed oligos to 500 µl 1 × NEB Buffer 3). 3. Ligation using Rapid Ligation Kit (Roche Diagnostics, Indianapolis, IN): Mix 2 µl of the annealed oligos with 2 µl vector and H2O (~5 ng) (oligos: vector, ~15:1). Add 1 µl DNA dilution buffer, mix well, briefly spin down. Add 5 µl ligation buffer and mix. Add 0.5 µl T4 DNA ligase, mix well, spin down briefly. Incubate 5 min at RT. 4. Transform into a suitable cloning bacterial strain using standard transformation protocols.
3.2.5
Cell Culture and Transfection
HeLa cells were maintained in DMEM supplemented with 10% (v/v) calf serum, 100 µg/ml penicillin, and 100 µg/ml streptomycin. For transfections, cells were grown in 6-well or 12-well plates and then switched into Opti-MEM I media before being transfected with 1 or 0.5 µg of DNA per well using LipofectAMINE Plus. Four hours post transfection, the media was replaced with fresh growth medium and the cells incubated for an additional 48 h (the time for collecting cells depends on the half-life of target genes).
3.2.6
Western Blotting
Twenty micrograms of total cell lysates were separated using 8% (w/v) SDS/PAGE, transferred to a nitrocellulose membrane, blocked with 1% casein, probed overnight with primary antibodies, washed, and incubated with secondary antibody conjugated to Alexa 680 or IRDye 800. Fluorescent signals were detected with an Odyssey infrared imaging system from LI-COR Biosciences – Biotechnology (Lincoln, NE, USA). Alexa 680 and IRdye 800-labeled secondary antibodies are scanned at 700 and 800 channels, respectively.
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3.3 3.3.1
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Results Generation of Vectors for Expressing Artificial miRNAs from a Synthetic Intron
Understanding the delicate chemistry of the RNAi machinery is perhaps the most important aspect of designing miRNA-based RNAi vectors. There are several key enzymes and complexes involved in the miRNA pathway, each recognizing a specific structural element within the RNA [3, 7]. The more recent miRNA vectors sandwiched artificial miRNA expression cassettes between an RNA polymerase II promoter and a fluorescent protein reporter gene [5, 12]. For example, the miR155 and miR30-based vectors coexpressed the miRNAs and EGFP segments as a combined exonic transcript [5, 12] (Fig. 3.1A). These vectors were designed to suppress the target gene expression by mature miRNAs and also provide an EGFP marker to indicate which cells had been successfully transfected. However, the EGFP signal that can be generated from these vectors arise only from the very low number of pri-miRNAs that are exported into the cytoplasm prior to Drosha cleavage (Fig. 3.1A). As a result of this flaw in the design, many cells, although transfected with the miRNA silencer, may not express a detectable level of the EGFP marker. Our group has generated a new miRNA expression strategy that is based on the endogeneous splicing mechanism of pri-miRNAs to overcome the original flaw in the miRNA vector design [6]. Instead of inserting a miRNA cassette into an exon, we have placed the miRNA-expressing cassette into a chimeric intron containing the 5′ intron donor site of the human β-globin gene and the branch and 3′ intron acceptor sites from an immunoglobulin gene (Fig. 3.1B). Two such vectors, pSM155 and pSM30, were derived from miR155 and miR30, respectively (Fig. 3.2). Based on our current understanding of miRNA processing, the new strategy would allow Drosha processing of the pri-miRNA to produce both a functional pre-miRNA and EGFP mRNA that can both be transported to the cytoplasm and perform their functions [3, 6, 7]. Indeed, these two vectors successfully downregulated expression of exogenously expressed luciferase and an endogenous phospholipase D2 (PLD2) gene while also generating a bright EGFP signal in transfected cells [6].
3.3.2
Cloning of miRNAs
The cloning of artificial miRNAs follows the common steps of cloning: oligo synthesis, vector preparation, oligo annealing, ligation, transformation, and characterization. Selection of miRNA oligos is based on the general guidelines for siRNA, which take into account miRNA stability and structure [9, 14]. An effective artificial miRNA must meet most of the following guidelines: overall low to medium G/C content (30–50%), low internal stability at the 5′ antisense strand, high internal stability at the 5′ sense strand, absence of internal repeats or palindromes, A-form
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Fig. 3.1 Strategies for miRNA-based RNAi vector. (A) Original miRNA-based vectors contained a miRNA-expressing cassette, such as miR155, sandwiched between a CMV promoter and EGFP as a combined exonic transcript. Upon processing, the EGFP gene segment is left uncapped and will be degraded quickly. Only a low number of unprocessed pri-mRNAs escape the nucleus and can be translated to generate the EGFP protein. Because expression of the EGFP marker is significantly decreased, it no longer serves as a reliable marker for miRNA-expressing cells. (B) The design of the new pSM155 vector. Placing the miRNA cassette into a synthetic intron is predicted to increase the reliability of the EGFP marker expression in miRNA-expressing cells since both functional miRNA and EGFP can be efficiently exported into the cytoplasm and expressed
3 RNA Interference Expression Vectors Based on miRNAs and RNA Splicing
Fig. 3.2 Plasmid maps for pSM155 and pSM30. These vectors are designed to express artificial miRNAs for RNAi experiments by incorporating the sequences of the human miR155 and miR30 miRNA precursors into a synthetic intron. The maps were generated using VectorNTI (Full sequences and digital files are available upon request)
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helix between miRNA and target mRNA, presence of an A at position 3 and 19 of sense strand, absence of a G or C at position 19 of sense strand, presence of a U at position 10 of sense strand, absence of a G at position 13 of sense strand, etc. To enhance specificity, a miRNA also needs to have minimal homology with non-target RNAs and avoid low-stringency sequences. A pair of oligos includes cohesive ends and a specific sequence matching target mRNA for each artificial miRNA (64 nucleotides for the pSM155-based system and 67 nucleotides for the pSM30-based system). For each gene, we usually test four to five target sequences and choose at least two constructs for our experiments. To simplify the cloning of artificial miRNAs without substantially altering the miRNA arm sequences, inverted BsmBI sites were placed internal to the arms of pSM30 and pSM155 (Fig. 3.3). A pair of oligonucleotide primers with appropriate 4 nt overhangs can be easily ligated to the cohesive sites of the vector generated by BsmB I digestion. Examples for the oligo sequences and their cloning in pSM155 and pSM30 are illustrated in Fig. 3.3. miR-203.1 miR-221/222 miR-200/429
Human FMR1 3 UTR
miR-148/152 miR-181
miR-124.2/506 miR-130/301 miR-23
miR-125/351 miR-141/200a
miR-9 miR-19
miR-205
miR-490
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Human APP 3 UTR
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2k (total predicted sites-91) NM_002024 3 UTR length:2248
miR-15/16/195/424/497 miR-153 miR-93.hd/291-3p/294/295/302/372/373/520 miR-101 miR-383 miR-17-5p/20/93.mr/106/519.d
miR-101
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miR-25/32/92/363/367 miR-194 miR-23 miR-194 miR-130/301 miR-101 miR-153
miR-19
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0.7k 0.8k 0.9k 1k 1.1k (total predicted sites-41) NM 000484 3 UTR length:1119
Human PSEN1 3 UTR
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miR-9
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Human LEPR 3 UTR miR-200b/429 1k
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Human NPY 3 UTR
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miR-33
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(total predicted sites-6) NM_ 000905 3 UTR length:172
Key:
1
0 Conservation
Site conserved in Human, Mouse, Rat, Dog,Chiken 8mer 7mer-m8 7mer-1a
Less conserved site 8mer 7mer-m8
7mer-1a
Fig. 3.3 Strategy for cloning specific artificial miRNA sequences into the targeting vectors. Two inverted BsmB I restriction sites are used for cloning a pair of oligos into pSM155 (A) and pSM30 (B) vectors. BsmB I digestion leaves the miRNA arms unchanged and generates two different cohesive ends into which a synthetic DNA duplex can be inserted to replace the original miR155 or miR30 sequences. The cloning of artificial miRNA sequences against luciferase (underlined) is shown as an example. The central black font indicates the loop region
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After ligation, the candidate recombinant clones need to be confirmed by restriction digestion and sequencing. The cloning efficiency of our current strategy is excellent and there is very low empty vector background. We often purify plasmids from two colonies for further characterization. For the pSM155 vector, the plasmid DNA is digested with Msc I. A linearized parental vector is 5,114 bp, and recombinant construct generates a 2,784 bp and a 2,368 bp fragment (Fig. 3.4A). For pSM30 vector, the plasmid DNA is digested with Xho I/Nhe I (Fig. 3.4B). The empty pSM30 vector is used as control. The recombinant clone and the vector release a 239 bp and 200 bp fragment, respectively. Finally, the candidate recombinant clones need to be confirmed by sequencing because the error rate of long oligos from commercial sources is often very high. Both pSM155 and pSM30 can also be used to express natural miRNAs, and thus label cells transfected with exogenously introduced natural miRNAs. A full-length miRNA can be amplified by PCR using a top primer with a 5′ flanking Sal I restriction site and a bottom primer with a 5′ flanking Nhe I, EcoR V or Mlu I restriction site. The PCR product digested with restriction enzymes is then cloned into pSM155 or pSM30 previously cut with the same enzymes (see maps in Fig. 3.2).
3.3.3
Conforming the Expression of EGFP Marker
As discussed above, directly linking the marker ORF to a miRNA-based artificial miRNA expression cassette as shown in Fig. 3.1A may lead to inefficient translation of the marker protein [3, 7]. We have shown efficient expression of the artificial miRNA and marker from a single RNA transcript in our vectors [6], suggesting that
Fig. 3.4 Characterization of the recombinant constructs. (A) Cutting of the parental vector with Msc I generates a 5,114 bp fragment (lane 2), whereas the recombinant pSM155 construct generates a 2,784 bp and a 2,368 bp fragments (lane 3). (B) Cutting of the pSM30 vector (lane 2) and the recombinant construct (lane 3) with Xho I/Nhe I releases a 200 and 239 bp fragment, respectively. Lane 1 in A and B shows the GeneRuler 1 kb DNA Ladder Plus
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RNA processing in our design is as predicted in Fig. 3.1B. We describe here our methods used to determine the expression of the EGFP marker in miRNA expression constructs generated from pSM155 and pSM30 vectors. To ensure that some miRNAs do not interfere with the expression of EGFP, the validation of efficient EGFP expression in the transfected cells is important before further functional analysis is done. Two methods can be used to evaluate EGFP expression: Western blotting or immunofluorescent microscopy. We prefer to use the latter since it allows us to compare the expression of EGFP with a second red fluorescent protein such as mCherry in the same cells. In summary, EGFP and mCherry fluorescent signals are compared in cells cotransfected with artificial miRNAs directed against genes of interest and pcDNA3.1-mCherry, which encodes a red fluorescent protein and serves as a marker for transfected cells. Such analysis was described earlier in our study to demonstrate that EGFP was efficiently expressed from the pSM155 and pSM30 vectors expressing the artificial miRNAs directed against firefly luciferase or PLD2, but poorly in the original miRNA expression vectors, pmiR155 and pmiR30 [6] (Fig. 3.5).
3.3.4
Determination of the Inhibition of Target Gene Expression
Knockdown efficiency is often determined by Western blotting. An irrelevant artificial miRNA construct such as that targeting luciferase can be used as a control. In some cases, immunofluorescent microscopy can also be used. If an antibody for the gene of interest is not available, reverse transcription-PCR is then the preferred method. Figure 3.6 illustrates PLD2 knockdown by Western blot analysis using the Odyssey Infrared Imaging System from LI-COR Biosciences – Biotechnology. One key to get reliable results is getting high transfection efficiency. We only perform Western blotting when more than 80–90% cells are transfected, as judged by the expression of EGFP. However, the functions of many genes have to be studied in particular cell types, which are sometimes hard to transfect. If these genes do not express or express at low levels in highly transfectable cell lines, determining the knockdown efficiency of each construct would be problematic. In this case, we usually determine which constructs are able to suppress the expression of exogenously expressed target genes. The cells are co-transfected with the miRNA constructs and the gene of interest tagged with an epi-tag such as Flag or Myc which can be detected by an antibody against the epi-tag using Western blotting. This method is very fast and reliable, especially when many constructs need to be tested. However, every candidate construct has to be tested for its ability in downregulating the expression of endogenous cognate genes before functional experiments are performed.
pre-miR-130b Chr 22 P13
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DGCR8 q11.21 q11.22 q11.23 q12.1 q12.2
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hsa-mir-33a
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Stimulating protein 1, ubiquitous zinc finger transcription factor Pleomorphic adenoma gene (PLAG) 1, a developmentally regulated C2H2 zinc finger protein
gtcagagggcaccctttccccccgggcagaggcccCgccccagccagcctgcattccaggtctcagatcc
Core promoter-binding protein (CPBP) with 3 Krueppel-type zinc fingers
q12.3
hsa-mir-658 hsa-mir-659
mature miR-130b
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hsa-mir-648 hsa-mir-185 hsa-mir-649 hsa-mir-301b hsa-mir-130b hsa-mir-650
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b C-allele
MYC-associated zinc finger protein related transcription factor EGR1, early growth response 1
G-allele Neural-restrictive-silencer-element
gtcagagggcaccctttccccccgggcagaggcccGgccccagccagcctgcattccaggtctcagatcc c
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Fig. 3.5 Determining the expression of EGFP marker proteins in artificial miRNA-expressing cells. miRNAs designed to target luciferase (luc) and PLD2 are used as examples. HeLa cells were cotransfected with pmiR155-luc, pmiR155-PLD2, pSM155-luc, or pSM155-PLD2, and pcDNA3.1/mCherry, which encodes a red fluorescent protein and is used to identify transfected cells. Both pmiR155 and pSM155 constructs contain an EGFP marker as illustrated in Fig. 3.1. Whereas EGFP is expressed in all cells expressing mCherry when the pSM155 vector is used, it is only expressed in a few cells when pmiR155 is used. Similar results were seen using the miR30-based vectors, pmiR30 and pSM30 (not shown here) (Modified and reproduced from [6]. With permission from Wiley-Blackwell Publishing Ltd.)
3 RNA Interference Expression Vectors Based on miRNAs and RNA Splicing
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Fig. 3.6 Determination of knockdown of PLD2 using Western blotting. HeLa cells were transfected with artificial miRNAs against luciferase (control) and PLD2 in pmiR155 and pSM155. Cell lysates were collected for Western blotting 2 days after transfection. PLD2 and α-tubulin were detected by a polyclonal antibody and a mouse monoclonal antibody, respectively, followed by goat anti-rabbit secondary IgG conjugated to Alexa 680 and goat anti-mouse conjugated to IRDye 800. Fluorescence was recorded using an Odyssey infrared imaging system from LI-COR Bioscience-Biotechnology (Lincoln, NE, USA) (Reproduced from [6]. With permission from Wiley-Blackwell Publishing Ltd.)
3.4
Discussion
We have shown that insertion of the miRNA-based artificial miRNA expression cassette into an intron significantly increased expression of the marker protein. A similar strategy for expressing the artificial miRNA from an intron was also recently reported by other groups [2, 8, 15]. In these studies, the miRNA expression cassettes were placed into the introns of an endogenous gene, i.e., the first intron of the human ubiquitin C gene [2, 15]. In our work, we utilized a synthetic intron to maximalize mRNA processing and directly compared the efficiencies of RNAi and marker gene expression in the original and our modified vectors [6]. Our results demonstrate that incorporation of an intronic strategy offers a modest, at best, improvement in the efficiency of RNAi yet generates a dramatic improvement in marker gene expression. This result suggests that Drosha processing of the pri-miRNA is relatively efficient even when the miRNA cassette is in an exon, however, most of the marker protein expression is lost through degradation of the resulting unstable mRNA that lacks a 5′ CAP structure (Fig. 3.1A). The success of vectors using a synthetic intron also indicates that the conserved sequences for mRNA splicing (5′ donor, branch, and 3′ acceptor sites) suffice for the efficient processing of pri-miRNAs. In summary, the miRNA expression vectors we describe here, pSM155 and pSM30, which are designed based on knowledge of miRNA and RNA splicing, provide a better approach to achieve efficient expression of both the RNAi cassette and the marker gene for transiently transfected cell experiments. The miRNA-based RNAi vectors also offer some technical advantages that may be useful in multiple applications. Dual shRNA and/or miRNA expression vectors can be prepared by subcloning a vector carrying the miRNA cassette for one miRNA into another vector carrying a different miRNA cassette (Fig. 3.7A).
3 RNA Interference Expression Vectors Based on miRNAs and RNA Splicing
Fig. 3.7 Some potential technical applications of pSM155 and pSM30 vectors. (A) Generation of constructs expressing two artificial miRNAs. Two constructs carrying two different miRNAs or shRNAs can be combined together to generate one construct containing both miRNA cassettes. To do so, construct 1 can be cut with restriction enzymes Xba I and Mlu I, and construct 2 with Nhe I and Mlu I. Because Xba I and Nhe I generate the same sticky ends, these two miRNA cassettes can be ligated into one vector using the standard ligation protocol. *Xba I (1793) site in pSM155 vector is methylated in Dam + E. coli strains and can’t be cut. Xba I cuts the plasmids purified from Dam + E. coli strains only at the Xba I (827) site. (B) Expression of miRNA and an antibiotic selection marker from the same mRNA transcript. An antibiotic selection marker can be cloned into the RNAi vector in place of EGFP. When generating stable cell lines, the antibiotic-resistant clones should all express miRNAs. (C) Performing RNAi and rescue experiments using a single construct. Rescue miRNA vectors can be generated within the same construct by replacing the EGFP cDNA for that of the gene of interest containing wobble mutations at the site targeted by the miRNA. The siRNA expressed from this construct is able to inhibit the expression of the mRNA from the endogenous locus but not that from the same construct
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Strategically placed restriction enzymes allow generation of one modified RNAi vector carrying two miRNAs. This can be a powerful technique with two major applications. In one case, both shRNAs can target different regions of the same target mRNA to increase silencing efficiency. In another case, the vector can comprise of two shRNAs that target two different genes for silencing. This application could then be used to silence related or opposing genes in one experiment. The latter application would be a beneficial technique especially for those studying multiple signal pathways as well as systems biology, since shRNA-expressing cells would be labeled even in tissues or whole animals. Expression of two miRNA cassettes that target different regions of the same gene can also avoid ineffectivity caused by selective mutations (or highly occurring mutations) of target sequences in some diseases. The ability to effectively express two synthetic miRNAs from a single transcript has been shown in a similar design [2]. The pSM155 and pSM30 vectors can also be used as a method to select true clones when used to generate stable cell lines expressing artificial miRNAs. For this case, the EGFP marker can be replaced by antibiotic selection markers (Fig. 3.7B). Since both miRNA and selection marker are expressed from a single mRNA, only cells expressing the artificial miRNA will be able to grow in growth media containing appropriate antibiotics. Another advantage of our vectors lies in the fact that RNAi and rescue experiments can be performed using the same vector. This would allow for more reliable and conclusive RNAi experiments. To do this, for example, the cDNA for the target mRNA can be cloned into the vector in place of or in addition to the EGFP gene (Fig. 3.7C). The cDNA should carry a wobble mutation so that the miRNA can no longer target it. This technique is useful to ensure the phenotype of the cell is due to direct silencing of the target gene and not a result of nonspecific targeting of the miRNA to other genes. Furthermore, this strategy can also be used to remove the mutated genes causing aberrant signaling in human diseases such as cancer, and replace them with their wild-type copies, which are often required to mediate normal physiological functions. Since the expression of miRNAs is driven by pol II promoters, the major therapeutic advantage of our vectors and other miRNA-based RNAi vectors is the ability to modify the vector for tissue specificity. Conjugated delivery methods for synthetic siRNAs can target only limited tissues and organs [4]. Introducing tissuespecific promoters into miRNA-based vectors will allow targeting to specific diseased tissues to reduce affecting normal cells and tissues. A promoter inducibly controlled by small molecules can also be adapted to drive miRNAs expression to avoid chronic toxicity of miRNA expression. Finally, while miRNA-based RNAi vectors including our design offer several advantages discussed above, it is not without its own limitations. One such limitation is that expression of miRNAs using retroviral vectors may suffer from low viral titers. As with miRNAs, viral RNA molecules are first processed and degraded by Drosha in the nuclei prior to moving to the cytoplasm. Only few unprocessed viral RNA can be transported to the cytoplasm as in Fig. 3.1A, resulting in low viral titer.
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In an intronic transcript (Fig. 3.1B), the miRNA expression cassette will be removed from mature viral RNAs, which will not packed into the retroviral particles. The second concern is that miRNA-based RNAi may not be as efficient as synthetic siRNAs and the traditional shRNAs driven by pol III promoters in some cell types and tissues. Maturation of siRNAs from the more complex miRNA structure requires at least one more processing step catalyzed by Drosha, which may eventually lead to generating less mature siRNAs. In fact, it has been reported that many miRNA primary transcripts are present at high levels but are not processed by the enzyme Drosha in early mouse development and human primary tumors [13]. This finding implies that we need to be more cautious in using miRNA-based vectors in some applications. Acknowledgements The authors thank Dr. Yoshiko Banno for the PLD2 antibody, Dr. Greg Hannon for the pSM2 vector, and Dr. Roger Y. Tsien for pRSET-B-mCherry. We also thank Dr. Michael Frohman for scientific discussion. This work was supported by a Scientist Development Grant from the American Heart Association (0430096 N) and research grants from National Institutes of Health (GM071475) to GD.
References 1. Brummelkamp, T.R., Bernards, R., and Agami, R. (2002). A system for stable expression of short interfering RNAs in mammalian cells. Science 296, 550–553. 2. Chung, K.H., Hart, C.C., Al-Bassam, S., Avery, A., Taylor, J., Patel, P.D., Vojtek, A.B., and Turner, D.L. (2006). Polycistronic RNA polymerase II expression vectors for RNA interference based on BIC/miR-155. Nucleic Acids Res 34, e53. 3. Cullen, B.R. (2004). Transcription and processing of human microRNA precursors. Mol Cell 16, 861–865. 4. de Fougerolles, A., Vornlocher, H.P., Maraganore, J., and Lieberman, J. (2007). Interfering with disease: a progress report on siRNA-based therapeutics. Nat Rev Drug Discov 6, 443–453. 5. Dickins, R.A., Hemann, M.T., Zilfou, J.T., Simpson, D.R., Ibarra, I., Hannon, G.J., and Lowe, S.W. (2005). Probing tumor phenotypes using stable and regulated synthetic microRNA precursors. Nat Genet 37, 1289–1295. 6. Du, G., Yonekubo, J., Zeng, Y., Osisami, M., and Frohman, M.A. (2006). Design of expression vectors for RNA interference based on miRNAs and RNA splicing. FEBS J 273, 5421–5427. 7. Kim, V.N. (2005). MicroRNA biogenesis: coordinated cropping and dicing. Nat Rev Mol Cell Biol 6, 376–385. 8. Lin, S.L., and Ying, S.Y. (2006). Gene silencing in vitro and in vivo using intronic microRNAs. Methods Mol Biol (Clifton, NJ) 342, 295–312. 9. Mittal, V. (2004). Improving the efficiency of RNA interference in mammals. Nat Rev Genet 5, 355–365. 10. Paddison, P.J., Caudy, A.A., Bernstein, E., Hannon, G.J., and Conklin, D.S. (2002). Short hairpin RNAs (shRNAs) induce sequence-specific silencing in mammalian cells. Genes Dev 16, 948–958. 11. Silva, J.M., Li, M.Z., Chang, K., Ge, W., Golding, M.C., Rickles, R.J., Siolas, D., Hu, G., Paddison, P.J., Schlabach, M.R., et al. (2005). Second-generation shRNA libraries covering the mouse and human genomes. Nat Genet 37, 1281–1288.
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12. Stegmeier, F., Hu, G., Rickles, R.J., Hannon, G.J., and Elledge, S.J. (2005). A lentiviral microRNA-based system for single-copy polymerase II-regulated RNA interference in mammalian cells. Proc Natl Acad Sci USA 102, 13212–13217. 13. Thomson, J.M., Newman, M., Parker, J.S., Morin-Kensicki, E.M., Wright, T., and Hammond, S.M. (2006). Extensive post-transcriptional regulation of microRNAs and its implications for cancer. Genes Dev 20, 2202–2207. 14. Ui-Tei, K., Naito, Y., Takahashi, F., Haraguchi, T., Ohki-Hamazaki, H., Juni, A., Ueda, R., and Saigo, K. (2004). Guidelines for the selection of highly effective siRNA sequences for mammalian and chick RNA interference. Nucleic Acids Res 32, 936–948. 15. Zhou, H., Xia, X.G., and Xu, Z. (2005). An RNA polymerase II construct synthesizes shorthairpin RNA with a quantitative indicator and mediates highly efficient RNAi. Nucleic Acids Res 33, e62.
Chapter 4
Recent Application of Intronic MicroRNA Agents in Cosmetics Shi-Lung Lin1*, David T.S. Wu2, and Shao-Yao Ying1
Abstract Utilization of gene silencing effectors, such as microRNA (miRNA) and small hairpin RNA (shRNA), provides a powerful new strategy for human skin care in vivo, particularly for hyperpigmentation treatment and aging prevention. For example, tyrosinase (Tyr), a melanocytic membrane-bound glycoprotein, is the rate-limiting enzyme critical for melanin (black pigment) biosynthesis in skins and hairs. There are over 54 native microRNA capable of targeting human tyrosinase for skin whitening and lightening. In this study, we have designed a mir-434-5p homologue and used it to successfully demonstrate the feasibility of miRNA-mediated skin whitening in vitro and in vivo. Under the same experimental condition in trials, Pol-II-directed intronic mir-434-5p expression did not cause any detectable sign of cytotoxicity, whereas siRNAs targeting the same sequence induced certain non-specific mRNA degradation as previously reported. Because the intronic miRNA-mediated gene silencing pathway is tightly regulated by multiple intracellular surveillance systems, including Pol-II transcription, RNA splicing, exosome digestion and NMD processing, the current findings underscore the fact that intronic miRNA agents, such as mir-434-5p homologues, are effective, target-specific and safe to be used for skin whitening without any overt cytotoxic effect. Given that the human skins also express a variety of native miRNAs, we may re-design these miRNAs based on their individual functions for skin care, which will provide significant insights into areas of opportunity for new cosmetic interventions. Keywords microRNA (miRNA), intronic microRNA (Id-miRNA), mir-434-5p, tyrosinase (Tyr), hyaluronidase (Hyal), RNA interference (RNAi), skin whitening, antiaging, cosmetics.
1
Department of Cell and Neurobiology, Keck School of Medicine, University of Southern California, 1333 San Pablo Street, BMT-302, Los Angeles, CA 90033, USA
2
Mello Biotech Ltd, Taipei, Taiwan, R.O.C.
* Corresponding author: Phone: 002-1-323-442-1658; Fax: 002-1-323-442-3466; E-mail:
[email protected] S.-Y. Ying (ed.) Current Perspectives in microRNAs (miRNA), © Springer Science + Business Media B.V. 2008
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Introduction
Prevention of hyperpigmentation (i.e. sun-burn) and aging is the key means for having healthy skins. However, many of the skin pigmentation and aging processes are associated with personal gene activities. For example, tyrosinase (Tyr), a melanocytic membrane-bound glycoprotein, is the rate-limiting enzyme critical for melanin (black pigment) biosynthesis in skins and hairs, while hyaluronidase (Hyal) often causes skin wrinkle by degrading subcutaneous hyaluronan (HA), the major anti-aging extracellular matrix in skins. Therefore, a good skin care can be achieved by suppressing these unwanted gene activities. Among a variety of currently available skin whitening and lightening products, many chemical and naturally extracted agents have been applied to inhibit tyrosinase function, using materials such as hormone-derived inhibitory oligopetides, hydroxytetronic acid derivatives, benzoyl compounds, hydroquinone compositions, alcohol diol and triol analogues, kojic acid derivatives, ascomycete-derived enzymes, and plant extracts. Although these cosmetic agents may work well in vitro, only a few of them, such as hydroquinone and its derivatives, are able to induce good hypopigmenting effects in clinical trials [25]. Nevertheless, all hydroquinone derivatives leading to a reactive quinone are putative cytotoxic agents. Thus, the gap between in-vitro and in-vivo studies suggests that innovative strategies are needed for validating their safety and efficacy. With the advance of recent RNA interference (RNAi) technologies, novel small RNA agents have been found to provide more potent effects in targeted gene suppression, including the utilization of double-stranded short interfering RNA (e.g. dsRNA/siRNA) [3, 4] and deoxyribonucleotidylated-RNA interfering molecules (e.g. D-RNAi) [12]. Conceivably, these small RNA agents may be used to develop new cosmetic designs and products for skin care. In principle, the RNAi mechanism elicits a post-transcriptional gene silencing (PTGS) phenomenon capable of inhibiting specific gene function with high potency at a few nanomolar dosage, which has been proven to be effective longer and much less toxic than conventional gene-knockout methods using antisense oligonucleotides or small molecule chemical inhibitors [12]. As reported in many previous studies [3, 6, 12, 14], the siRNA-induced gene silencing effects may last over one week, while the D-RNAi effects can even sustain up to one month after one treatment. These siRNA/D-RNAi agents evoke a series of intracellular sequence-specific mRNA degradation and/or translational suppression processes, affecting all highly homologous gene transcripts, namely co-suppression. It has been observed that such co-suppression results from the generation of small RNA products (21–25 nucleotide bases) by the enzymatic activities of RNaseIII endoribonucleases (Dicer) and/or RNA-directed RNA polymerases (RdRp) on aberrant RNA templates, which are usually the derivatives of foreign transgenes or viral genomes [3, 6, 12].
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Limitations of SiRNA/ShRNA-Based Gene Silencing Agents
Although the modern RNAi technologies may offer a new avenue for suppressing unwanted gene function in skins, the applications thereof have not been demonstrated to work constantly and safely in higher vertebrates, including fish, avian, mammal and human. For example, almost all of the current siRNA agents are based on a double-stranded RNA (dsRNA) conformation, which has been shown to cause interferon-mediated non-specific RNA degradation in vertebrates [3, 26]. Such an interferon-mediated cytotoxic response reduces the target specificity of siRNAinduced gene silencing effects and often results in global RNA degradation in vertebrate cells. Particularly in mammalian cells, it has been noted that the RNAi effects are disturbed when the siRNA/dsRNA size is longer than 25 base-pairs (bp) [3]. Transfection of siRNA or small hairpin RNA (shRNA) sized less than 25 bp may not completely overcome such a problem, because both [24] and [15] have reported that the high dosage of siRNAs and shRNAs (such as >250 nM in human T cells) is able to cause strong cytotoxic effects similar to those of long dsRNAs. This toxicity is due to their double-stranded RNA conformation, which activates the interferon-mediated non-specific RNA degradation and programmed cell death through the activation of cellular PKR and 2–5A signaling pathways. It is well known that interferon-activated protein kinase PKR can trigger cell apoptosis, while the activation of interferon-induced 2′,5′-oligoadenylate synthetase (2–5A) system leads to extensive cleavage of single-stranded RNAs, such as mRNAs [26]. Both PKR and 2-5A systems contain dsRNA-binding motifs, which possess high affinity to the double-stranded RNA conformation. Further, the most difficult problem is that these small siRNA/shRNA agents are not stable enough to be maintained at an optimal dose in vivo due to the abundant RNase activities in higher vertebrates [1]. As the RNAi effects are naturally caused by the production of small RNA products (21–25 nucleotide bases) from a transcriptional template derived from foreign transgenes or viral genomes [6, 12], the recent utilization of Pol-III-directed siRNA/ shRNA expression vectors has shown to offer relatively stable RNAi efficacy in vivo [27]. Although previous studies [9, 20, 22] using such a vector-based siRNA approach have succeeded in maintaining constant gene silencing effects, their strategies fail to focus the RNAi effects on a targeted cell or tissue population because of the ubiquitous existence of type III RNA polymerase (Pol-III) activity. Pol-III promoters, such as U6 and H1, are activated in almost all cell types, making tissue-specific gene silencing impossible. Moreover, because the leaky read-through activity of Pol-III transcription often occurs on a short DNA template in the absence of pr oper termination, large RNA products longer than desired 25 bp can be synthesized and cause unexpected interferon cytotoxicity [8, 23]. Such a problem can also result from the competitive conflict between the Pol-III promoter and another vector promoter (i.e. LTR and CMV promoters). Furthermore, it is recently noted that high
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siRNA/shRNA concentrations generated by the Pol-III-directed RNAi systems can over-saturate the cellular native microRNA (miRNA) pathway and thus cause global miRNA inhibition and cell death [7]. These disadvantages discourage the use of Pol-III-based RNAi vector systems in health care. In order to improve the delivery stability, targeting specificity and safety aspects of modern RNAi technologies for healthy skin care, a better transduction and maintenance strategy is highly desired.
4.3
Intronic MicroRNA-Mediated RNAi Mechanism
Research based on gene transcript (e.g. mRNA), an assembly of protein-coding exons, is fully described throughout the literature, taking the fate of spliced non-coding introns to be completely digested for granted [21]. Is it true that the intron portion of a gene is destined to be a genetic waste without function or there is a function for it, however, which has not yet been discovered? Recently, this misconception was corrected by the observation of intronic microRNA (miRNA) [13, 29, 30]. Intronic miRNA is a new class of small single-stranded regulatory RNAs derived from the gene introns, which are spliced out of the precursor messenger RNA (pre-mRNA) of the encoding gene and further processed into small mature miRNAs. MiRNA is usually about 18–27 nucleotides (nt) in length and is capable of either directly degrading its messenger RNA (mRNA) target or suppressing the protein translation of its targeted mRNA, depending on the complementarity between the miRNA and its target. In this way, the intronic miRNA is functionally similar to previously described siRNA/shRNA, but differs from them in the requirement of intracellular type II RNA polymerase (Pol-II) transcription and RNA splicing processes for its biogenesis [13]. Also, because introns naturally contain multiple translational stop codons for recognition by the intracellular nonsense-mediated decay (NMD) system [11, 31], most of the unstructured intron sequences can be quickly degraded after RNA splicing to prevent excessive accumulation, which is toxic to the cells. It has been measured that approximately 10–30% of a spliced intron is preserved after the exosome and NMD digestion in cytoplasm with a relatively long half-life, indicating the cellular origin of native intronic miRNAs [2]. Natural intronic miRNA biogenesis relies on the coupled interaction between nascent Pol-II-mediated pre-mRNA transcription and intron splicing/excision (Fig. 4.1), occurring within certain nuclear regions proximal to genomic perichromatin fibrils [5, 14]. In eukaryotes, protein-coding gene transcripts, such as mRNAs, are produced by type-II RNA polymerases (Pol-II). The transcription of a genomic gene generates precursor messenger RNA (pre-mRNA), which contains four major parts including 5′-untranslated region (UTR), protein-coding exon, non-coding intron and 3′-UTR. Broadly speaking, both 5′- and 3′-UTR can be seen as a kind of intron extension. Introns occupy the largest proportion of non-coding sequences in the pre-mRNA. Each intron can be ranged up to 30 or so kilo-bases and is required to be excised out of the pre-mRNA content before mRNA maturation. This process of pre-mRNA excision and intron removal is called RNA splicing, which is executed by intracellular
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Fig. 4.1 Biogenesis of native intronic microRNA (miRNA). Intronic miRNA is co-transcribed with precursor messenger RNA (pre-mRNA) by Pol-II and cleaved out of the pre-mRNA by RNA splicing, while the ligated exons become a mature messenger RNA (mRNA) for protein synthesis. The spliced intronic miRNA with a high secondary structure (i.e. hairpin and/or stem-loop) is further processed into mature miRNA capable of triggering RNAi-related gene silencing effects
spliceosomes. After RNA splicing, some of the intron-derived RNA fragments are further processed to form microRNA (miRNA) derivative molecules, which can effectively silence their targeted genes, respectively, through an RNA interference (RNAi)-like mechanism, while exons of the pre-mRNA are ligated together to form a mature mRNA for protein synthesis.
4.4
Differences Between miRNA and siRNA Biogenesis Pathways
We have demonstrated that effective mature miRNAs can be generated from the introns of vertebrate genes, of which the biogenetic process is different from those of siRNA and intergenic miRNA [13, 16]. To demonstrate their differences, Fig. 4.2 shows the comparison of native biogenesis and RNAi mechanisms among siRNA, intergenic (exonic) miRNA and intronic miRNA. Presumably, siRNA is formed by two perfectly complementary RNAs transcribed by two reversely positioned promoters from one DNA template, then hybridized and further processing into 20–25 bp duplexes by RNaseIII endoribonucleases, namely Dicer. Different from this siRNA model, the biogenesis of intergenic miRNA, e.g. lin-4 and let-7, involves a long non-coding precursor RNA transcript (pri-miRNA), which is directly transcribed from a Pol-II or Pol-III RNA promoter, whereas intronic miRNA is co-transcribed with its encoding gene by only Pol-II and released after
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Fig. 4.2 Comparison of biogenesis and RNAi mechanisms among siRNA, intergenic (exonic) miRNA and intronic miRNA. SiRNA is likely formed by two perfectly complementary RNAs transcribed from two different promoters (remains to be determined) and further processing into 19–22 bp duplexes by RNaseIII-familial endonucleases, Dicer. The biogenesis of intergenic (exonic) miRNA, e.g. lin-4 and let-7, involves a long transcript precursor (pri-miRNA), which is probably generated by a Pol-II or Pol-III RNA promoter, whereas intronic miRNA is mainly transcribed by the Pol-II promoter of its encoded gene and co-expressed in the intron region of the gene transcript (pre-mRNA). After pre-mRNA splicing, the spliced intron functions as a primiRNA for intronic miRNA generation. In the nucleus, the pri-miRNA is excised by either Drosha-like RNases (intergenic miRNA) or spliceosomal components (intronic miRNA) to form a hairpin-like pre-miRNA template and then exported to cytoplasm for further processing by Dicer* to form mature miRNAs. The Dicers for siRNA and miRNA pathways are different. For instance, some exosome and NMD components are likely involved in the process of intronic miRNA maturation. All three small regulatory RNAs are finally incorporated into a RNA-induced silencing complex (RISC), which contains either strand of siRNA or the single-strand of miRNA. The effect of miRNA is considered to be more specific and less adverse than that of siRNA because only one strand is involved. On the other hand, siRNAs primarily trigger mRNA degradation, whereas miRNAs can induce either mRNA degradation or suppression of protein synthesis depending on the sequence complementarity to the target gene transcripts
RNA splicing as a spliced intron. The spliced intron is then served as a pri-miRNA for processing into an intronic precursor miRNA (pre-miRNA) or a multipre-miRNA cluster. In the cell nucleus, the pri-miRNA is further excised by either Drosha-like RNases (for intergenic miRNA) or spliceosomal components (for intronic miRNA) to form a hairpin-like stem-loop precursor or a cluster of multiple
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stem-loop structures, termed pre-miRNA, and then exported to cytoplasm for final processing into mature miRNA by a miRNA-associated Dicer (Dicer*). Subsequently, all three small regulatory RNAs are incorporated into a RNA-induced silencing complex (RISC), which contains either strand of siRNA or the mature strand of miRNA. The Dicers and RISCs for siRNA and miRNA pathways are known to be different [28]. For example, some enzymes of the nonsense-mediated decay (NMD) system may play the role of Dicer* in intronic miRNA maturation. As a result, the effect of miRNA is generally more specific and less adverse than that of siRNA because only one strand is involved. On the other hand, siRNAs primarily trigger mRNA degradation, whereas miRNAs can induce either mRNA degradation or suppression of protein synthesis, or both, depending on the sequence complementarity to their targeted gene transcripts. Because the intronic miRNA pathway is well coordinated by multiple intracellularly regulatory systems, including Pol-II transcription, RNA splicing and NMD processing, the gene silencing effects of intronic miRNAs are considered to be effective, specific and safe [19].
4.5
Development of miRNA-Based Gene Silencing Agents
Based on the intronic RNA splicing and processing mechanisms (Figs. 3A, B), we designed and developed a Pol-II-mediated recombinant gene expression system containing at least a splicing-competent intron, namely SpRNAi, which is able to inhibit the function of a unwanted gene with high complementarity to the intron sequence. The SpRNAi is co-transcribed with the precursor mRNA (pre-mRNA) of the recombinant gene by Pol-II RNA polymerases (P) and cleaved out of the premRNA by RNA splicing. Subsequently, the spliced SpRNAi was further processed into mature gene silencing agents, such as shRNA and miRNA, capable of triggering RNAi-related gene silencing. After intron removal, the exons of the recombinant gene transcript are linked together to form a mature mRNA molecule for translational synthesis of a marker or functional protein. As shown in Fig. 4.3A, the essential components of the SpRNAi intron include several consensus nucleotide elements, consisting of a 5′-splice site, a branch-point motif (BrP), a poly-pyrimidine tract (PPT), and a 3′-splice site. In addition, a hairpin RNA-like pre-miRNA sequence is inserted inside the SpRNAi intron located between the 5′-splice site and the branch-point motif (BrP). This portion of the intron would normally form a lariat structure during RNA splicing and processing. We have observed that spliceosomal U2 and U6 snRNPs, both helicases, are involved in the unwinding and excision of the lariat RNA fragment into pre-miRNA; however, the detailed processing remains to be elucidated. Further, the 3′-end of the SpRNAi construct contains a multiple translational stop codon region (T codon) in order to increase the accuracy of intronic RNA splicing and NMD processing. When presented in a cytoplasmic mRNA, this T codon would signal the activation of the nonsense-mediated decay (NMD) pathway to degrade any unstructured RNA accumulation in the cell. However, the highly secondary structured hairpin RNA and
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Fig. 4.3 Structural composition of the SpRNAi-incorporated recombinant RGFP gene (SpRNAiRGFP) in an expression-competent vector (A), and the strategy (B) of using this composition to generate man-made microRNA, mimicking the biogenesis of the natural intronic miRNA. In vivo tests of an SpRNAi-RGFP expression composition directed against green EGFP in fish show an over 85% knockdown effect specifically on the targeted EGFP gene expression, as determined by Western blot analysis (C). The intron-derived anti-EGFP microRNA and its spliced precursor can be observed on a 1% formaldehyde agarose gel electrophoresis after Northern blot analysis (D)
pre-miRNA insert will be preserved for further Dicer cleavage, so as to form mature siRNA and miRNA, respectively. Moreover, for intracellular expression, we manually incorporate the SpRNAi construct in the DraII restriction site of a red fluorescent protein (RGFP) gene isolated from mutated chromoproteins of the coral reef Heteractis crispa, so as to form a recombinant SpRNAi-RGFP gene. The cleavage of RGFP at its 208th nucleotide site by the restriction enzyme DraII generates an AG– GN nucleotide break with three recessing nucleotides in each end, which will form 5′- and 3′-splice sites respectively after the SpRNAi insertion. Because this intronic insertion disrupts the structure of a functional RGFP protein, which can be recovered by intron splicing, we can determine the release of intronic shRNA/miRNA and RGFP-mRNA maturation through the appearance of red RGFP around the affected
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cells. The RGFP gene also provides multiple exonic splicing enhancers (ESEs) to increase RNA splicing accuracy and efficiency. In this intronic miRNA expression system (Fig. 4.3B), we provides a genetic engineering method for using synthetic RNA splicing and processing elements, such as 5′-splice site, branch-point motif (BrP), poly-pyrimidine tract (PPT), and 3′-splice site, to form an artificial SpRNAi intron containing at least a desired RNA insert for antisense RNA, small hairpin RNA (shRNA) and/or microRNA (miRNA) production. A DNA synthesizer can chemically produce and link these elements. Alternatively, the linkage of these elements can be achieved by enzymatic restriction and ligation. The intron so obtained can be used directly for transfection into cells of interest or further incorporated into a cellular gene for co-expression along with the gene transcript (i.e. pre-mRNA) by Pol-II. During RNA splicing and mRNA maturation, the desired RNA insert will be excised and released by intracellular spliceosome, exosome and NMD mechanisms and then triggers a desired gene silencing effect on specific gene transcripts with high complementarity to the inserted RNA sequence, while the exons of the recombinant gene transcript are linked together to form mature mRNA for expression of a desirable gene function, such as translation of a reporter or marker protein selected from the group of red/green fluorescent protein (RGFP/EGFP), luciferase, lac-Z, and their derivative homologues. The presence of the reporter/marker protein is useful for locating the production of the inserted shRNA/miRNA molecules in affected cells, facilitating the identification of the desired gene silencing/RNAi effects. In accordance with the biogenesis of intronic miRNA, mature mRNA formed by the linkage of exons can also be useful in conventional gene therapy to replace impaired or missing gene function, or to increase specific gene expression. Alternatively, this method provides novel compositions and means for inducing cellular production of gene silencing molecules through intronic RNA splicing and processing mechanisms to elicit either antisense-mediated gene knockout or RNA interference (RNAi) effects, which are useful for inhibiting targeted gene function. The intron-derived gene silencing molecules so obtained may include antisense RNA, ribozyme, short temporary RNA (stRNA), double-stranded RNA (dsRNA), small interfering RNA (siRNA), tiny non-coding RNA (tncRNA), short hairpin RNA (shRNA), microRNA (miRNA), and RNAi-associated precursor RNA constructs (pri-/pre-miRNA). The use of these intronic RNA-derived gene silencing agents is a powerful tool for targeting and silencing unwanted genes selected from the group consisting of pathogenic transgenes, viral genes, mutant genes, oncogenes, disease-related small RNA genes and any other types of protein-coding as well as non-coding genes. Using this novel Pol-II-mediated SpRNAi-RGFP expression system, we have successfully generated mature shRNA and miRNA molecules with full gene silencing capacity in human prostate cancer LNCaP, human cervical cancer HeLa and rat neuronal stem HCN-A94–2 cells [17] as well as in zebrafish, chicken and mouse in vivo [18]. We have tested different pre-miRNA insert constructs targeting against green EGFP and other cellular gene expression in zebrafish and various human cell lines, and have learned that effective gene silencing miRNAs are
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derived from the 5′-proximity of the intron sequence between the 5′-splice site and the branching point. As shown in Fig. 4.3C, a strong gene silencing effect occurs only in the transfection of anti-EGFP pre-miRNA insert (lane 4), whereas no effect can be detected in those of other inserts indicated by lanes from left to right: 1, blank vector control (Ctl); 2, pre-miRNA insert targeting HIV-p24 (mock); 3, antisense EGFP insert without the hairpin loop structure (anti); and 5, reverse premiRNA sequence which is completely complementary to the anti-EGFP pre-miRNA (miR*). No effect was detected on off-target genes, such as marker RGFP and house-keeping β-actin, suggesting that such intronic miRNA-mediated RNA interference (RNAi) is highly target-specific. To further confirm the role of RNA splicing in this intronic RNAi effect, we have also tested three different SpRNAi-RGFP expression systems as shown in Fig. 4.3D by lanes from left to right: (1) vector expressing intron-free RGFP without any pre-miRNA insert; (2) vector expressing RGFP with an intronic anti-EGFP pre-miRNA insert; and (3) vector similar to the 2 construct but with a defective 5′-splice site in the SpRNAi intron. As a result of this, Northern bolt analysis shows that mature miRNA is released only from the spliced intron of the vector 2 construct, which is exactly identical to the SpRNAi vector construct with the anti-EGFP pre-miRNA insert in the Fig. 4.3C, indicating the requirement of cellular RNA splicing for intronic miRNA biogenesis.
4.6
Optimization of Intronic miRNA Designs
After the above understanding, we have further determined the optimal structural design of the pre-miRNA inserts for inducing maximal gene silencing effects and learned that a strong structural bias exists in the cellular selection of a mature miRNA strand during assembly of the RNAi effector, the RNA-induced gene silencing complex (RISC) [16]. RISC is a protein–RNA complex that directs either target gene transcript degradation or translational repression through the RNAi mechanism. Formation of siRNA duplexes plays a key role in assembly of the siRNA-associated RISC. The two strands of the siRNA duplex are functionally asymmetric, but assembly into the RISC complex is preferential for only one strand. Such preference is determined by the thermodynamic stability of each 5′-end base-pairing in the strand. Based on this siRNA model, the formation of miRNA and its complementary miRNA (miRNA*) duplexes was thought to be an essential step in the assembly of miRNA-associated RISC. If this were true, no functional bias would be observed in the stem-loop structure of a pre-miRNA. Nevertheless, we observed that the stemloop orientation of the intronic pre-miRNA is involved in the strand selection of a mature miRNA for RISC assembly in zebrafish. To find the correct miRNA structures for RISC assembly, we have constructed two different intronic pre-miRNA-inserted SpRNAi-RGFP expression vectors containing a pair of symmetric pre-miRNA constructs, respectively, namely miRNA*-stemloop-miRNA [1] and miRNA-stemloop-miRNA* [2], as shown in Fig. 4.4A. Both pre-miRNAs contain the same double-stranded stem-arm structure,
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which is directed against the EGFP nucleotide 280–302 sequence. In definition here, miRNA refers the exactly complete sequence of a mature microRNA, while miRNA* refers the reverse nucleotide sequence complementary to the mature microRNA sequence. After liposomal transfection of these miRNA-expressing SpRNAi-RGFP vectors (60 µg each) into two-week-old zebrafish larvae for 24 hours [16], we have isolated the zebrafish small RNAs using mirVana miRNA isolation columns (Ambion, Austin, TX) and then precipitated all the potential miRNAs matched to the targeted EGFP region by latex beads containing the target sequence. After sequencing, one effective miRNA identity, miR-EGFP(280–302),
Fig. 4.4 Different designs of intronic RNA inserts in an SpRNAi-RGFP construct for effective microRNA biogenesis. Gene silencing of a targeted green fluorescent protein (EGFP) expression in Tg(actin-GAL4:UAS-gfp) zebrafish demonstrates the asymmetric preference of RISC assembly between the transfection of [1] 5′-miRNA*-stemloop-miRNA-3′ and that of [2] 5′-miRNAstemloop-miRNA*-3′ hairpin RNA structures, respectively (A). In vivo gene silencing is only observed in the transfection of the [2] pre-miRNA construct, but not the [1] construct. Since the color combination of EGFP and RGFP displays more red than green (as shown in deep orange), the expression level of target EGFP (green) is significantly reduced in the [2] pre-miRNA transfection, while vector indicator RGFP (red) is evenly present in all vector transfections (B). Western blot analysis of the EGFP protein levels confirms the specific silencing result of the [2] premiRNA transfection (C). No detectable gene silencing is observed in fish with other treatments, such as liposome only (Lipo), empty vector without any insert (Vctr), and siRNA (siR)
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is identified to be active in the transfections of the 5′-miRNA-stemloop-miRNA*-3′ construct [2], as shown in Fig. 4.4A (gray-shading sequences). Since the mature miRNA is detected only in the zebrafish transfected by the [2] construct, the miRNA-associated RISC must preferably interact with the construct [2] rather than the [1] pre-miRNA, demonstrating the existence of a structural bias for intronic miRNA–RISC assembly. In this experiment, we use an actin-promoter-driven Tg(UAS:gfp) strain zebrafish, namely Tg(actin-GAL4:UAS-gfp), which constitutively express a green fluorescent EGFP protein in almost all cell types of the fish body. As shown in Fig. 4.4B, transfection of the SpRNAi-RGFP vector in these zebrafish silences the targeted EGFP and co-expresses a red fluorescent marker protein RGFP, serving as a positive indicator for intronic miRNA generation in the affected cells. The gene silencing effect in the gastrointestinal (GI) tract is somehow lower than other tissues, probably due to the high RNase activity in this region. Based on further Western blot analysis (Fig. 4.4C), the indicator RGFP protein expression is detected in both of the fish transfected with either 5′-miRNA*-stemloop-miRNA-3′ [1] or 5′-miRNA-stemloop-miRNA*-3′ [2] pre-miRNA, whereas gene silencing of the target EGFP expression (green) only occurs in the fish transfected with the [2] premiRNA construct, confirming the result of Fig. 4.4B. Because thermostability of the 5′-end stem-arm of both pre-miRNA constructs is the same, we conclude that the stem-loop of the intronic pre-miRNA is involved in the strand selection of a mature miRNA sequence during RISC assembly. Given that the cleavage site of Dicer in the stem-arm is known to determine the strand selection of mature miRNA [10], the stem-loop of an intronic pre-miRNA may function as a determinant for the recognition of the special cleavage site. In this early design, because the over sizes of many native pre-miRNA stemloop structures cannot fit in the SpRNAi-RGFP expression vector for efficient expression, we must use a short tRNAmet loop (i.e. 5′-(A/U)UCCAAGGGGG-3′) to replace the native pre-miRNA loops. The tRNAmet loop has been shown to efficiently facilitate the export of designed miRNAs from nucleus to cytoplasm through the same Ran-GTP and Exportin-5 transporting mechanisms [16]. Later, we use a pair of manually improved pre-mir-302 loops (i.e. 5′-GCTAAGCCAGGC3′ and 5′-GCCTGGCTTAGC-3′), which provide the same nuclear export efficiency as the native pre-miRNAs but not interfere with the tRNA exportation. The design of these new pre-miRNA loops is based on a mimicking modification of short stem-loops of mir-302s, which are highly expressed in embryonic stem cells but not in other differentiated tissue cells. Thus, the use of these man-made pre-miRNA loops will not interfere with the native miRNA pathway in the adult human body. For different pre-miRNA generation, because the intronic insertion site of the recombinant SpRNAi-RGFP gene is flanked with a PvuI and an MluI restriction site at its 5′- and 3′-ends, respectively, the primary intronic insert can be easily removed and replaced by various gene-specific pre-miRNA inserts (e.g. anti-EGFP and anti-Tyr pre-miRNA) possessing matched cohesive ends. By changing the pre-miRNA inserts directed against different gene transcripts, this intronic miRNA generation
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system can be served as a powerful tool for inducing targeted gene silencing in vitro and in vivo. For confirming the correct insert size, the pre-miRNA-inserted SpRNAirGFP gene (10 ng) can be amplified by a polymerase chain reaction (PCR) with a pair of oligonucleotide primers (i.e. 5′-CTCGAGCATG GTGAGCGGCC TGCTGAA-3′ and 5′-TCTAGAAGTT GGCCTTCTCG GGCAGGT-3′) for 25 cycles at 94 °C, 52 °C and then 70 °C each for one minute. The resulting PCR products are fractionated on a 2% agarose gel, and then extracted and purified by gel extraction kit (Qiagen, Valencia, CA) for sequencing confirmation.
4.7
Evaluation of Natural Anti-tyrosinase miRNA Agents
We adopt the proof-of-principle design of the Pol-II-mediated SpRNAi-RGFP expression system and use it for developing new cosmetic products for skin care. In this new approach, we apply skins a non-naturally occurring intron capable of being processed into hairpin-like precursor microRNA (pre-miRNA) molecules by the skin cells and thus inducing specific gene silencing effects on epidermal pigment-related genes and/or aging-causing genes. In this case, the RNA splicing- and processing-generated gene silencing molecule is the hairpin-like pre-miRNA insert located within the intron area of the recombinant gene and is capable of silencing a targeted gene, such as tyrosinase (Tyr), hyaluronidase (Hyal), hyaluronan receptors CD44 and CD168, and other pigmentation-related and/or aging-related genes and oncogenes. Alternatively, such a pre-miRNA insert can also be artificially incorporated into the intron region of a cellular gene in the skin. In general, this kind of intronic insertion technology includes plasmid-like transgene transfection, homologous recombination, transposon delivery, jumping gene integration and retroviral infection. In the present design, the recombinant SpRNAi-RGFP gene expresses an intronic insert construct reminiscent of a hairpin-like pre-mRNA structure. The recombinant gene is consisted of two major different parts: exon and intron. The exon part is ligated after RNA splicing to form a functional mRNA and protein for identification of the intronic RNA release, while the intron part is spliced out of the recombinant gene transcript and further processed into a desired intronic RNA molecule, serving as a gene silencing effector, including antisense RNA, miRNA, shRNA, siRNA, dsRNA and their precursors (i.e. pre-miRNA and piRNA). These desired intronic RNA molecules may comprise a hairpin-like stem-loop structure containing a sequence motif homologous to 5′-GCTAAGCCAG GC-3′ or 5′-GCCTGGCTTA GC-3′, which facilitates not only accurate excision of the desired RNA molecule out of the intron but also nuclear exportation of the desired RNA molecule to the cell cytoplasm. Also, the stem-arms of these intron-derived RNA molecules contain homology or complementarity, or both, to a targeted gene or a coding sequence of the targeted gene transcript. The homologous or complementary sequences of the desired RNA molecules are sized from about 18 to about 27 nucleotide bases. The homology and/or complementarity rate of the desired
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intronic RNA molecule to the targeted gene sequence is ranged from about 30–100%, more preferably 35–49%, for a desired hairpin-like intronic RNA and 90–100% for a linear intronic RNA molecule. In addition, the 5′-end of the non-naturally occurring intron contains a donor splice site homologous to 5′-GTAAGAGK-3′ motifs, while its 3′-end is a acceptor splice site that is homologous to 5′-GWKSCYRCAG-3′ motifs. Moreover, a branch point sequence is located between the 5′- and 3′-splice sites, containing homology to 5′-TACTWAY-3′ motifs. The adenosine “A” nucleotide of the branch-point sequence forms a part of (2′–5′)-linked lariat intron RNA by cellular (2′-5′)-oligoadenylate synthetases and spliceosomes in almost all spliceosomal introns. Furthermore, a poly-pyrimidine tract is closely located between the branch-point and 3′-splice site, containing a high T or C content oligonucleotide sequence homologous to either 5′-(TY)m(C/−)(T)nS(C/−)-3′ or 5′(TC)nNCTAG(G/−)-3′ motifs. The symbols of “m” and “n” indicate multiple repeats ≥1; most preferably, the m number is equal to 1~3 and the n number is equal to 7~12. The symbol “–” refers an empty nucleotide in the sequence. There are also some linker nucleotide sequences for the connection of all these intron components. In definition, the symbol W refers to an adenine (A) or thymine (T)/uracil (U), the symbol K refers to a guanine (G) or thymine (T)/uracil (U), the symbol S refers to a cytosine (C) or guanine (G), the symbol Y refers to a cytosine (C) or thymine (T)/uracil (U), the symbol R refers to an adenine (A) or guanine (G), and the symbol N refers to an adenine (A), cytosine (C), guanine (G) or thymine (T)/uracil (U).” Based on the above design, we have tested an optimized SpRNAi-RGFP gene construct expressing either anti-Tyr or anti-Hyal pre-miRNA directed against the unwanted pigmentation-related gene Tyr or aging-related gene Hyal in mouse skins (Fig. 4.3A). These pre-miRNAs target a highly conserved region (>98% homology) in both human and mouse Tyr and Hyal genes, respectively. In nature, there are 54 native miRNAs capable of targeting human tyrosinase (Tyr; 2082 bp) for pigmentation gene silencing, including mir-1, mir-15a, mir-16, mir-31, mir-101, mir-129, mir-137, mir-143, mir-154, mir-194, mir-195, mir-196b, mir-200b, mir200c, mir-206, mir-208, mir-214, mir-221, mir-222, mir-292-3p, mir-299-3p, mir-326, mir-328, mir-381, mir-409-5p, mir-434-5p, mir-450, mir-451, mir-452, mir-464, mir-466, mir-488, mir-490, mir-501, mir-522, mir-552, mir-553, mir-570, mir-571, mir-582, mir-600, mir-619, mir-624, mir-625, mir-633, mir-634, mir-690, mir-697, mir-704, mir-714, mir-759, mir-761, mir-768-5p, and mir-804. According to the miRNA-target database of the miRBase:: Sequences program (http://microrna.sanger.ac.uk), all these anti-Tyr miRNAs are directed against a region within the first 300 nucleotides of the Tyr gene transcript (NCBI accession number NM000372). Moreover, there are 9 native miRNAs capable of targeting hyaluronidase (Hyal; 2518 bp; NCBI accession number NM007312) for aging gene silencing, including mir-197, mir-349, mir-434-5p, mir-549, mir-605, mir-618, mir-647, mir-680, mir-702, and mir-763. In these native miRNAs, the mir-434-5p is the only one targets both Tyr and Hyal genes in human and also it is one of the most efficient miRNAs targeting the least off-target genes other
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than Tyr and Hyal. However, because almost all native miRNAs have several to over fifty targets and they tend to bind with some of the target genes more strongly than others, the use of these native miRNAs is likely not specific and safe enough for the skin care purpose. To test the feasibility of miRNA-mediated skin whitening, we have utilized the SpRNAi-RGFP expression system to express native pre-mir-434-5p in mouse skin. As shown in Fig. 4.5, patched albino (white) skins of melanin-knockdown mice (W-9 black) can be created by a succession of intra-cutaneous (i.c.) injections of the pre-mir-434-5p expression vector (50 µg) directed against tyrosinase (Tyr) for four days (total 200 µg). The Tyr, a type-I membrane protein and copper-containing enzyme, catalyzes the critical and rate-limiting step of tyrosine hydroxylation in the biosynthesis of melanin (black pigment) in skins and hairs. Starting from about two
Fig. 4.5 Depigmentational effects of RNAi-mediated tyrosinase (Tyr) gene silencing on mouse skins and hairs, indicating the feasibility of targeted gene knockdown in epidermal tissues using subcutaneous transfection of the recombinant SpRNAi-RGFP gene vector expressing a native mir434 pre-miRNA insert. Transfection of this mir-434-5p expression construct induces a strong and specific gene silencing effect on Tyr but not house-keeping GAPDH expression, whereas that of a U6 promoter-based siRNA expression vector against the same Tyr target sequence triggers nonspecific RNA degradation of both Tyr and GAPDH gene transcripts. Because Tyr plays an essential role in melanin (black pigment) production, the successful Tyr gene silencing can be observed by a significant loss of the black color in mouse skins and hairs. The circles indicate the location of i.c. injections. Small windows show the Northern blotting of Tyr mRNA knockdown in local hair follicles, confirming the effectiveness of the mir-434-mediated gene silencing effect
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weeks after the first i.c. injection, we observe that skin and hair pigments was significantly lost only in the pre-miRNA transfections. On the contrary, the blank control and the Pol-III (U6)-directed siRNA transfections present no significant effect. Northern blot analysis using mRNAs isolated from the hair follicles of the pre-mir-434-5p transfections show a 76.1% ± 5.3% reduction of Tyr expression two days post-transfection, whereas mild, non-specific degradation of random gene transcripts is detected in the siRNA-transfected skins (seen from the smearing patterns of both house-keeping control GAPDH and targeted Tyr mRNAs). Since [7] have reported that high siRNA/shRNA concentrations generated by the Pol-IIIdirected RNAi systems can over-saturate the cellular microRNA pathway and cause global miRNA dysregulation, this result indicates that the siRNA pathway is incompatible with the native miRNA pathway in skin tissues. Thus, the use of miRNA will likely provide a more effective, compatible and safe means for skin care. However, because the native mir-434-5p also targets five other cellular genes for silencing, including TRPS1, PITX1, LCOR, LYPLA2 and Hyal, the off-target effect of this native pre-mir-434-5p transfection remains to be determined.
4.8 Re-design of mir-434-5p for Skin Whitening Use in Human In order to improve the target-specificity and safety of anti-Tyr miRNA agents, we have re-designed the seed sequence of the mir-434-5p to form a highly matched region binding to either Tyr nucleotides 3–25 (namely miR-Tyr) or Hyal nucleotides 459–482 (namely miR-Hyal). The pre-miRNA insert sequence for Tyr gene silencing (pre-miR-Tyr) is 5′-GTCCGATCGT CGCCCTACTC TATTGCCTAA GCCGCTAAGC CAGGCGGCTT AGGCAATAGA GTAGGGCCGA CGCGTCAT-3′, which will form a hairpin-like RNA after splicing and will be further processed into a mature miR-Tyr microRNA (miRNA) sequence containing or homologous to 5′GCCCTACTCT ATTGCCTAAG CC-3′. Alternatively, the pre-miRNA insert for Hyal gene silencing (pre-miR-Hyal) is 5′-GTCCGATCGT CAGCTAGACA GTCAGGGTTT GAAGCTAAGC CAGGCTTCAA ACCCTGACTG TCTAGCTCGA CGCGTCAT-3′, which will form a different kind of hairpin-like RNA after splicing and will be further processed into a mature miR-Hyal miRNA sequence containing or homologous to 5′-AGCTAGACAG TCAGGGTTTG AA-3′. Although both pre-miR-Tyr and pre-miR-Hyal constructs are re-designed based on the same mir-434-5p backbone and mir-302 stem-loop, the mature miR-Tyr and miR-Hyal so obtained are totally different from each other. As shown in Fig. 4.6, the transfective expressions of miR-Tyr and miR-Hyal in mouse skins specifically knock down the targeted Tyr (reduction >90%) and Hyal genes (reduction > 67%), respectively, without any crossover off-target effect. The expression levels of mature miR-Tyr and miR-Hyal microRNAs are directly measured by Northern blot analysis, while the knockdown rates of the targeted Tyr and Hyal gene products (proteins) are determined by Western blot analysis.
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Fig. 4.6 Improvement of Tyr gene silencing using a man-made anti-Tyr pre-miRNA (miR-Tyr) insert expressed by the recombinant SpRNAi-RGFP gene vector in mouse skins, showing a more specific and less off-target gene silencing effect on the targeted tyrosinase (Tyr) gene. Neither offtarget (hyaluronidase) nor house-keeping (ß-actin) genes are affected by the transfection of this man-made intronic miR-Tyr microRNA
After understanding the optimized gene silencing effects of the re-designed miRTyr and miR-Hyal miRNAs in mice, we continue to test their efficacy, target specificity and safety in human skins. For efficient vector transfection into the human epidermal cell layers, a 1 µg/ml SpRNAi-RGFP vector solution is made by mixing 100 µg of the purified SpRNAi-RGFP vector in 1 ml of autoclaved ddH2O with 99 ml of 100% DNase-free glycerin (or called glycerol). DNase-free glycerin is used to encapsulate miR-Tyr for deep skin delivery and cell membrane penetration. This forms the major ingredient base for skin whitening and lightening products. Based on this, more other cosmetic ingredients may be added to increase the color, fragrance, effectiveness and/or stability of the final cosmetic products. As shown in Fig. 4.7A, Asian male arm skins treated with 2 ml of this major ingredient base expressing the aforementioned miR-Tyr (right site) versus empty SpRNAi-RGFP vector without any miRNA insert (glycerin control, left site) are compared. The result of skin whitening (loss of the black pigment–melanin) by the miR-Tyr treatment can be clearly observed in three days after two single treatments per day, as shown in this figure.
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Fig. 4.7 Human trial results of the improved anti-Tyr pre-miRNA (miR-Tyr) insert expressed by the recombinant SpRNAi-RGFP gene vector, identical to the Fig. 4.6 approach but in the human arm skins (A) and primary skin cell cultures (B and C), showing an over 50% knockdown rate in tyrosinase (Tyr) expression as determined by Western blot analysis
Then, primary skin cell culture is obtained by trypsin-dissociated skin explants from the tested donor with personal consent. The SpRNAi-RGFP vector transfection (final 6 µg/ml) in the primary skin culture is performed using a FuGene liposomal reagent (Roche Biochemicals, Indianapolis, IN), as described previously [13, 2006a]. Figure 4.7B shows that Western blot analyses of the loss of the targeted tyrosinase proteins and its substrate melanin are biostatistically significant (p > 0.001). The reduction amounts of tyrosinase proteins and its substrate melanin in skins is proportional to the treated concentrations of the miR-Tyr expression vector, indicating the positive correlation between the increase of the miR-Tyr treatment and the loss of the targeted tyrosinase proteins and its substrate melanin. No effect is found in other treatments, such as an empty SpRNAi-RGFP vector without any miRNA insert (glycerin) and an SpRNAi-RGFP vector expressing an anti-EGFP pre-miRNA insert (miR-gfp). At the concentration of 1 µg/ml of the miR-Tyr expression vector transfection, the optimal Tyr gene silencing rate is approximately 55–60% for tyrosinases and 30–45% for melanin, while the expression of non-target house-keeping control ß-actin is not affected by the miR-Tyr treatment, indicating the high target-specificity of this man-made microRNA molecule. Figure 4.7C further shows that the skin melanin levels are significantly reduced as shown in bright-field (BF) photographs of the primary skin cell culture (upper
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panels), while melanin (black dots around the cell nuclei) is highly expressed in the normal skin cells without the miR-Tyr treatment (i.e. blank and glycerin only). The miR-Tyr-treated skin cells present very limited melanin accumulation, demonstrating an effective skin-whitening effect in vivo. In regard to this loss of skin melanin, the targeted tyrosinase expression is concurrently reduced in the miR-Tyr-treated skin cells, as determined by immunocytochemical (ICC) staining analysis (Fig. 4.7C, lower panels). Therefore, based on these results, the re-designed miR-Tyr microRNA can be used to knock down the tyrosinase expression and successfully blocks melanin production in the human skins in vivo.
4.9
Microarray Analyses of Target Specificity and Safety
After establishing the gene silencing efficacy of the miR-Tyr in human skins, we use gene microarray analysis (Human GeneChip U133A&B arrays, Affymetrix, Santa Clara, CA) to assess the changes of approximately 32,668 human gene expression in the above miR-Tyr-transfected versus non-treated primary skin cell cultures, showing a much more target-specific and less offtarget gene silencing effect than the use of native mir-434. Total RNAs from each tested cell culture is isolated using RNeasy spin columns (Qiagen, CA). As shown in Fig. 4.8A (left), the result of microarray analysis in non-treated (miR–) versus miR-Tyr-transfected (miR+) primary skin cell cultures shows that there are only two genes altered more than 1.5 fold (>50% change of gene expression), including the targeted tyrosinase (Tyr) and its associated TRP1 gene (Fig. 4.8B), indicating that the miR-Tyr-mediated gene silencing effect is highly specific to the targeted Tyr. Furthermore, no gene related to either cytotoxicity or interferon-mediated PKR/2-5A pathways is affected, suggesting that this gene silencing effect is safe for skin care treatments. We have also used Northern blot analysis to compare and assess the gene expression levels of these microarray-identified genes (Fig. 4.8C), confirming the results of Figs. 4.8A, B. In further comparison with the result of the native mir-434-5p transfection (Fig. 4.8A, right), the correlation coefficiency (CC) rate clearly indicates that a high 99.8% population of the 32,668 tested human genes remains to be unchanged in the miR-Tyr-transfected (miR+) cells, while a low 77.6% CC rate is found in the mir-434-5p-transfected cells. This means that the expression patterns of at most only 65 cellular genes are altered by the redesigned miR-Tyr transfection, whereas those of over 7,317 genes may be changed by the native mir-434-5p transfection. Because it is a well-known fact that almost all native microRNAs (miRNAs) target multiple cellular genes due to their mismatched stem-arms, our present study demonstrates that the redesign of these stem-arm regions is required for the safe use of these miRNAs in target-specific gene silencing applications.
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Fig. 4.8 Gene microarray analysis (Affymetrix human GeneChip U133A&B, CA) of altered gene expression in the above human primary skin cell cultures with or without anti-Tyr pre-miRNA transfection, showing a much more target-specific and less off-target gene silencing effect than the use of native microRNAs, such as mir-434-5p
4.10
Conclusion
In sum, utilization of intronic hairpin-like microRNA (miRNA) expression provides a powerful new strategy for human skin care in vivo, particularly for hyperpigmentation treatment and aging prevention. Under the same treatment in animal trials, Pol-II-directed intronic miRNA expression does not cause any detectable cytotoxicity, whereas Pol-III-directed siRNAs induced non-specific mRNA degradation as previously reported [15, 24]. This underscores the fact that the intronic miRNA agent is effective, target-specific and safe in vivo. Because the intronic miRNAmediated gene silencing pathway is regulated by multiple intracellular surveillance systems, including Pol-II transcription, RNA splicing, exosome digestion and NMD processing, the gene silencing of intronic miRNA is considered to be the most effective, specific and safe approach among all three currently known RNAi
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pathways. Advantageously, using this intronic miRNA expression strategy, many cosmetic applications can be designed and developed for skin care, offering more long-term effectiveness, better target-specificity and higher safety in skin gene manipulation, which prevents the unspecific off-target cytotoxicity as commonly seen in the conventional siRNA methods.
References 1. Brantl S. (2002). Antisense-RNA regulation and RNA interference. Biochimica et Biophysica Acta 1575: 15–25. 2. Clement JQ, Qian L, Kaplinsky N, Wilkinson MF. (1999). The stability and fate of a spliced intron from vertebrate cells. RNA 5: 206–220. 3. Elbashir SM, Harborth J, Lendeckel W, Yalcin A, Weber K, Tuschl T. (2001). Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells. Nature 411: 494–498. 4. Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, Mello CC. (1998). Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391: 806–811. 5. Ghosh S, Garcia-Blanco MA. (2000). Coupled in vitro synthesis and splicing of RNA polymerase II transcripts. RNA 6: 1325–1334. 6. Grant SR. (1999). Dissecting the mechanisms of posttranscriptional gene silencing: divide and conquer. Cell 96: 303–306. 7. Grimm D, Streetz KL, Jopling CL, et al. (2006). Fatality in mice due to oversaturation of cellular microRNA/short hairpin RNA pathways. Nature 441: 537–541. 8. Gunnery S, Ma Y, Mathews MB. (1999). Termination sequence requirements vary among genes transcribed by RNA polymerase III. J Mol Biol 286: 745–757. 9. Lee NS, Dohjima T, Bauer G, et al. (2002). Expression of small interfering RNAs targeted against HIV-1 rev transcripts in human cells. Nat Biotechnol 20: 500–505. 10. Lee Y, Ahn C, Han J, et al. (2003). The nuclear RNase III Drosha initiates microRNA processing. Nature 425: 415–419. 11. Lewis BP, Green RE, Brenner SE. (2003). Evidence for the widespread coupling of alternative splicing and nonsense-mediated mRNA decay in humans. Proc Natl Acad Sci USA 100: 189–192. 12. Lin SL, Ying SY. (2001). D-RNAi (messenger RNA-antisense DNA interference) as a novel defense system against cancer and viral infections. Curr Cancer Drug Targets 1: 241–247. 13. Lin SL, Chang D, Wu DY, Ying SY. (2003). A novel RNA splicing-mediated gene silencing mechanism potential for genome evolution. Biochem Biophys Res Commun 310: 754–760. 14. Lin SL, Ying SY. (2004a). Novel RNAi therapy – intron-derived microRNA drugs. Drug Des Rev 1: 247–255. 15. Lin SL, Ying SY. (2004b). Combinational therapy for HIV-1 eradication and vaccination. Int J. Oncol 24: 81–88. 16. Lin SL, Chang D, Ying SY (2005). Asymmetry of intronic pre-microRNA structures in functional RISC assembly. Gene 356: 32–38. 17. Lin SL, Ying SY. (2006a). Gene silencing in vitro and in vivo using intronic microRNAs. Methods Mol Biol 342: 295–312. 18. Lin SL, Chang SJE, Ying SY. (2006b). Transgene-like animal model using intronic microRNAs. Methods Mol Biol 342: 321–334. 19. Lin SL, Kim H, Ying SY. (2008). Intron-mediated RNA interference and microRNA (miRNA). Front Biosci 13: 2216–2230.
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20. Miyagishi M, Taira K. (2002). U6 promoter-driven siRNAs with four uridine 3′ overhangs efficiently suppress targeted gene expression in mammalian cells. Nat Biotechnol 20: 497–500. 21. Nott A, Meislin SH, Moore MJ. (2003). A quantitative analysis of intron effects on mammalian gene expression. RNA 9: 607–617. 22. Paul CP, Good PD, Winer I, Engelke DR. (2002). Effective expression of small interfering RNA in human cells. Nat Biotechnol 20: 505–508. 23. Schramm L, Hernandez N. (2002). Recruitment of RNA polymerase III to its target promoters. Genes Dev 16: 2593–2620. 24. Sledz, CA, Holko M, de Veer MJ, Silverman RH, Williams BR. (2003). Activation of the interferon system by short-interfering RNAs. Nat Cell Biol 5: 834–839. 25. Solano F, Briganti S, Picardo M, Ghanem G. (2006). Hypopigmenting agents: an updated review on biological, chemical and clinical aspects. Pigment Cell Res 19: 550–571. 26. Stark GR, Kerr IM, Williams BR, Silverman RH, Schreiber RD. (1998). How cells respond to interferons. Annu Rev Biochem 67: 227–264. 27. Tuschl T. (2002). Expanding small RNA interference. Nat Biotechnol 20: 446–448. 28. Tang G. (2005). siRNA and miRNA: an insight into RISCs. Trends Biochem Sci 30: 106–114. 29. Ying SY, Lin, SL. (2004). Intron-derived microRNAs–fine tuning of gene functions. Gene 342: 25–28. 30. Ying SY, Lin SL. (2005). Intronic microRNAs. Biochem Biophys Res Commun 326: 515–520. 31. Zhang G, Taneja KL, Singer RH, Green MR. (1994). Localization of pre-mRNA splicing in mammalian nuclei. Nature 372: 809–812.
Chapter 5
MicroRNA Profiling in CNS Tissue Using Microarrays Reuben Saba1,2 and Stephanie A. Booth1,2*
Abstract MicroRNAs (miRNAs) are important regulators of gene expression in virtually all eukaryotic cell types including the diverse cell types found in the CNS. They are involved in repressing gene expression by complementary hybridization to cognate protein-coding mRNAs. The likely involvement of miRNAs in disease processes requires both accurate detection and expression analysis strategies. In comparison to conventional methodologies to study miRNA expression, microarrays offer an advantage in terms of throughput, sensitivity and specificity. Although microarrays are almost routinely used in laboratories for the analysis of mRNA, the small size of miRNAs presents challenges for their analysis in terms of probe design, target labeling and hybridization conditions. We discuss these issues in this chapter as well as highlighting the emerging perspectives in this field. Keywords miRNA, central nervous system, microarray, oligonucleotide linker, probe design, direct-labelling, indirect-labelling, normalization, qRT-PCR, laser capture microdissection
5.1
Introduction to miRNAs
MicroRNAs (miRNAs) are ~18–25 nt long post-transcriptional regulators of gene expression in both plants and animals [7]. MiRNAs are genome encoded and are derived from a much larger transcript called a pri-miRNA. Pri-miRNAs are processed in the nucleus by the enzyme Drosha to generate ~70 nt long, stem-loop transcripts known as pre-miRNAs. These are exported through a nuclear-pore complex into the cytoplasm and subsequently processed by the enzyme Dicer to generate mature miRNAs. Mature miRNAs are incorporated into an RNA-induced silencing complex 1
Department of Medical Microbiology, University of Manitoba, Winnipeg, MB, R3E 0W3, Canada
2
Prion Diseases Program, National Microbiology Laboratory, Public Health Agency of Canada, Winnipeg, MB, R3E 3R2, Canada * Corresponding author: E-mail:
[email protected] S.-Y. Ying (ed.) Current Perspectives in microRNAs (miRNA), © Springer Science + Business Media B.V. 2008
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(RISC), which either suppresses the translation and/or promotes the degradation of cognate protein-coding mRNA molecule(s) through complementary hybridization to the 3′-UTR. Among vertebrates, miRNAs are only partially complementary to their gene targets and only nucleotides 2–7 from the 5′-end of the miRNA, a region often referred to as the “seed sequence”, is predicted to bind to the target mRNA [40]. This imperfect complimentarity between the miRNA and its target site on the mRNA means that it is possible for any given miRNA to regulate hundreds of potential targets [34, 41]. It has been suggested that up to ~30% or more of the protein coding genes in the vertebrate genome are under miRNA regulation [40].
5.2
MiRNAs in the CNS
MicroRNAs have been implicated in several important biological functions in the CNS including neurogenesis [71], dendrite formation [59], brain morphogenesis [23], and silencing of non-neuronal transcripts [16, 64, 72]. One of the most abundant miRNAs identified in the adult mammalian brain is miR-124a which can account for up to 48% of the total miRNAs content of the brain [37]. Fascinatingly, this miRNA has been shown to play a putative role in modulating cell lineage fate. In one study, injection of miR-124a into HeLa cells (human carcinoma cell line) resulted in alterations in expression of over 100 different mRNA transcripts to generate a gene expression profile resembling that of a mature neuron [43]. Possibly, the binding motif of miR-124a, which is one of the most prevalent miRNA recognition elements (MREs) in the 3′-UTR of mammalian transcripts, is an important regulatory motif for the expression of neuronal genes and the maintenance of neuronal identity [76]. Due to the abundance of miRNAs in the CNS, as well as their regulatory and pleiotrophic properties, the involvement of these molecules in neurological disease processes is not surprising. The evidence for miRNA dysfunction in CNS disorders has been steadily accumulating including roles in Tourette’s syndrome [1], DiGeorge syndrome [38], fragile X mental retardation [30] and schizophrenia [26, 53]. More recently, evidence for the role of miRNA deregulation in neurodegenerative diseases including Alzheimer’s [47] and Parkinson’s [32] was also reported. In the latter, miR-133b was shown to work in an autoregulatory feedback loop with the transcription factor PitX3 to promote the maturation and survival of dopaminergic neurons in the brain. In patients with Parkinson’s disease and also in mouse models where there is a deficiency of dopaminergic neurons there is decreased expression of miR-133b [32]. Moreover, the deregulation of miRNAs in tumours of the CNS has been extensively studied (for a relevant review see [21]). Despite the fact that miRNAs have only recently been detected and characterized in the CNS, there exists a substantial amount of experimental evidence for their importance in the proper development and maintenance of this tissue. Furthermore, emerging evidence linking miRNA expression to dysfunctions of the CNS, advocates the targeting of these molecules as potential points of therapeutic intervention.
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Transcriptional Profiling of miRNAs Using Microarrays
The adaptation of genomic technologies to the study of miRNAs will ensure a rapid explosion in knowledge in this area over the next few years. Interestingly, in most publications that have laid down the groundwork for miRNA microarrays, the principal tissue studied was from the CNS. In this chapter, we describe the current trends, challenges and opportunities in the field of miRNA profiling using microarrays and other emerging technologies. We have examined a number of aspects of this methodology such as choice of array platform, the design of probes, miRNA isolation/enrichment options, target labelling strategies, hybridization conditions, and data processing. In several instances, novel strategies are presented that may be readily applied to the analysis of miRNAs from CNS tissues. Microarrays consist of addressable complimentary immobilized probes of DNA, RNA, or protein on a planar solid support, usually a glass slide [58]. The number of probes can vary from a few hundred in macro-projects to tens of thousands in large scale micro-projects. In this chapter, the word ‘probe’ refers to the reporter sequence, which is placed at a particular position on the microarray as it interrogates the sample for the presence of its reverse complement. The word ‘target’ refers to the molecule being interrogated in the sample. The probes on a microarray are arranged so that their specific location is known and can be referenced later on. Most of the current array experiments utilize fluorescent-tags to label targets that are subsequently detected by laser confocal microscopy. A quantitative inference of the abundance of the hybridized target from the original sample is determined from the intensity of the fluorescent signal on the array. Specifically, the relationship between the intensity of the signal to that of the abundance of the target is proportional.
5.3.1
MiRNA Array Platforms
The type of microarray platform utilized for profiling miRNAs plays a central role on the experimental design and type of analysis that can be performed. The user must choose between either custom or commercially available microarrays. Since commercial miRNA microarrays have only become available very recently, and the identification of miRNAs is still in progress, the use of custom manufactured microarrays is often the preferred option. The earliest prototype microarrays used for profiling miRNAs from CNS tissue were on filter or nylon membranes spotted with oligonucleotide sequences for mature miRNAs in either the sense [63] or antisense [35, 67] orientation. The predominant choice of microarray platform at present is planar glass; although the emerging use of liquid-phase bead-based microarrays is an interesting innovation [5] and will be discussed later in the chapter. The choices of immobilization chemistry for the attachment of probes to glass are wide-ranging. The most common are chemical coatings on the glass surface, such as aldehyde, epoxy, or poly-L-lysine,
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which interact either covalently or non-covalently with the DNA probe. A number of novel slide surfaces also show promise for the attachment of miRNA probes. The use of slides with three-dimensional surface matrices such CodeLink activated slides (GE Health Care, Piscataway, NJ, USA), Genopal® slides (Mitsubishi Rayon Co., Ltd., Yokohama, Japan) and slides that rely on evanescent resonator (ER) technology (UnaxisAG, Balzers, Liechtenstein) are emerging in the field. CodeLink Activated Slides are coated with a proprietary 3-D surface chemistry comprised of a long-chain, hydrophilic polymer containing amine-reactive groups. This polymer is covalently crosslinked to itself and to the surface of the slide. The crosslinked polymer, combined with end-point attachment, orients the immobilized DNA, and holds it upright from the surface of the slide. This combination means that the DNA is more readily available for hybridization and eliminates the need for stilts to hold up the capture probe. Additionally, the hydrophilic nature of the polymer provides a passive effect once the DNA has been immobilized. The overall result is a substantially lower background. A similar 3-D surface is used for miRNA probes spotted on Genopal® slides. In this instance it is plastic hollow fibers arranged in a block like fashion on the slide surface, initially cut from a large block of resin hardened fibers, which provides the 3-D space for attaching the probes [28]. Planar glass slides that employ Evanescent Resonance (ER) technology have greater sensitivity than conventional microarray slides due to their unique optical features [11] and have recently been introduced into miRNA research [9]. The distinguishing feature of these slides is a uniformly corrugated surface coated with a highly refractive index material, such as Ta2O5, that reduces background fluorescence and abnormal reflection patterns which may contribute to fluorescence cross talk. The overall effect of the ER platform is the improvement in the sensitivity of microarray detection which is imperative for low abundance miRNAs.
5.3.2
Probe Design
5.3.2.1
Probe Sequence
An invaluable resource for probe design is the freely available miRBase (http:// microrna.sanger.ac.uk) formerly known as the Sanger Registry [25]. This database houses the most up-to-date collection of miRNA sequences of both the pre-miRNAs (~60–70 nt) and mature miRNAs. It is possible to design microarrays to interrogate either pre-miRNAs or the mature species. Profiling the primary transcript (hairpin precursor) may provide valuable complementary information, however, most studies seek to understand the regulatory properties of mature miRNAs on protein expression, and so detection of mature miRNAs is generally the method of choice. Several strategies have been proposed to minimize cross-hybridization between closely related miRNAs on an array platform. In a recent publication it was shown that capture probes for miRNAs that have ~18–19 consecutive identical nucleotides show cross-hybridzation in comparison to probes for miRNAs that have less than
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18–19 consecutive identical nucleotides [67]. Therefore, it was suggested that probes should only be designed for miRNAs that have less than 18 consecutive identical bases. For example, within the let-7 family, probes should only be designed for let-7b, 7d, 7e, 7f, 7g, and 7i since they satisfy the criteria. However, probes for let-7a or let-7c would show cross-reactivity. In this case it may be possible to use other methods of identifying miRNAs that differ by a few nucleotides, such as qRT-PCR. In an alternate strategy, the large range of Tm values across all known miRNAs was curtailed in order to minimize the cross reactivity among closely related miRNAs. This was done by trimming the probes in a successive and alternating fashion from both the 3′- and 5′-ends, 1 nt at a time, until the Tm range between the probes was minimal [24]. Additional weight was given to trimming at the 5′-end of the probes which contains the highly conserved seed region so as to preserve the more variable 3′-end for better discrimination between closely related miRNAs.
5.3.2.2
Probe Immobilization via ‘Linker’ Addition
Determining optimal probe length for targets that are only ~18–25 nt long offers a unique challenge in the design of miRNA capture probes. Longer probe sequences offer more sensitivity with compromised specificity [54, 55], whereas short oligonucleotides provide greater specificity at the expense of sensitivity. Specificity of the probe also appears to be affected by steric hinderance caused by the proximity of the array surface. Nucleotide mismatches near the end of the probe that is tethered to the slide, can influence the specificity of a given probe to a greater extent than if the mismatch is towards the free end of the probe [5, 57]. The addition of ‘linker’ sequences to the probe overcomes these obstacles by holding the capture sequence away from the slide surface, thus reducing steric hinderence in the hybridization step, and allowing for a short, specific capture sequence to be used. The directional orientation of the nucleotide probes with respect to their binding to the microarray is not an issue in the design of probes which can be linked by either their 3′- or 5′-ends. Often, the technique used to generate the labeled target dictates the orientation of the probes and vice versa. A number of linkers, both nucleotide and chemical, have been described. These include using the pre-miRNA sequence (in which the miRNA capture sequence is embedded within the miRNA precursor sequence) [5], random ‘words’ that are absent in genomes [5], poly(T) tracts [57], and most often 5′-amino (6-carbon) linkers have been used when printing onto amine-coated slides (for example CodeLink slides) [8, 45, 48]. Another chemical linker that shows promise for the immobilization of miRNA capture probes is the addition of 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC). EDC cross linking likely proceeds via a 5′-terminal phosphate to form a phosphoramidate bond between the RNA and the nylon membrane. To date, EDC has only been used to immobilize size fractionated RNA on nylon membranes for Northern blot analysis of miRNAs [52]. In this case, EDC immobilized RNA was shown to be more amenable to hybridization with target, than probes covalently linked to the membrane by other means such as UV cross-linking.
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Chemical Modification of Probes
A number of probe modifications, independent from the addition of a ‘linker’, have been found to positively influence the specificity and sensitivity of target detection. Most of these probe modifications involve the incorporation, during synthesis, of nucleotide analogs that demonstrate more favourable hybridization characteristic than standard DNA-based probes. The most promising is the incorporation of locked nucleic acid (LNA) monomers into the probe sequence [14, 50, 69]. LNA nucleotides contain a methylene bridge that connects the 2′-oxygen of the ribose with the 4′-carbon [10, 69]. This bridge effectively locks the furnose ring in the sugar phosphate backbone and thereby reduces the conformational flexibility of the ribose. This results in an increase in the melting temperature (Tm) of the hybrid by +1−8 °C per LNA monomer for DNA hybrids and +2−10 °C per monomer in RNA hybrids [69], which in turn increases the affinity between probe and target, improves mismatch discrimination, and increases metabolic stability. One major outcome is that by adding varying amounts of LNA to array probes it is possible to design microarray capture probes with uniform Tm. In unmodified probes, the Tms can range between 45–74 °C thus compromising probe specificity and sensitivity during hybridization [14]. LNA modified probes have been used to improve the detection of CNS specific/enriched miRNAs by Northern blotting [69, 70], in situ detection [33, 50, 51] and by microarrays [11]. Another miRNA capture probe modification that has been investigated is the substitution at the 2′-postion of the ribose by O-(2-methoxyethyl) (MOE) side chain [9]. The addition of this moiety was shown to increase the affinity and specificity of binding to native RNA [44]. However, this was accompanied by a rather high false positive rate for the resulting microarray that was attributed to subjecting all probes on the array to identical hybridization temperatures; unlike LNA monomers which can be used to design a uniform Tm for the probes on an array. Though other promising oligonucleotide analogs exist, such as peptide nucleic acid (PNA), phosphoramidate DNA, hexitol nucleic acid (HNA) and morpholino based oligomers, they have not yet been tested in the design of capture probes for miRNA microarrays.
5.3.2.4
Probes for Use as Positive and Negative Controls
In miRNA microarray analysis, various control probes (positive, negative, and mismatch probes) serve to validate the sensitivity and specificity of the methodology. The most common positive controls used for analysis are other abundant small RNA species that are isolated alongside the miRNA from a biological sample. These include transfer RNAs (tRNAs), ribosomal RNAs (such as 5S RNA, 5.8S RNA), and U6 RNA species. The possibility of using these small RNA species as positive controls is only feasible when size fractionated mature RNAs are not used as starting material, as these control RNAs are significantly longer than the mature miRNA. If fractionated mature miRNAs are used then the only choice of positive control may be
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synthetic small RNAs spiked into the target; complementary probes to the spikes must be spotted on the array. To date, miRNA(s) that are ubiquitously expressed across all tissue types have not been identified, however, a number of miRNAs are known to be expressed at high levels in a tissue specific manner. In the case of CNS tissues, miR-124a may be an ideal positive control as this miRNA has been identified to be abundant in the majority of miRNA microarray analysis of this tissue [4, 5, 8, 9, 28, 35, 48, 57, 62, 77]. It is worth mentioning, however, that miR-124a has been recently shown to be present in abundance in the spinal cord [67], heart [67], and pancreas [6] suggesting that miRNAs may not be absolutely tissue specific. The most commonly utilized negative controls in vertebrate miRNA microarray research are plant miRNAs, as these miRNAs are predicted to be absent in the animal kingdom [57]. The main concern with using known animal miRNAs as negative controls is the extensive tissue tropism that is shown by animal miRNAs [60]. Another negative control that has been described is a trimer consisting of a random stretch of sequences (NHG-sequences; 10mer) that is very rare in the human genome [5]. Mismatch probes fulfill a further role in the determination of probe specificity; this is especially important in the validation of miRNA arrays as these molecules are often closely related, some differing by only a single nucleotide. Probes containing mismatches are often spotted alongside miRNA capture probes for at least a selection of miRNA probes on an array. Hybridization to mismatch probes has revealed that these probes generally tolerate 1–3 nucleotide differences between probe and target sequences. The extent of tolerance to mismatches, however, depends upon the location along the length of the probe. If the mismatch is in close proximity to the end of the probe that is tethered to the surface of the platform, then it is tolerated to a greater extent than if it was at the furthest end [5, 57, 67]. Tolerance, in this case, may be due to the limited accessibility of the wash buffer, during post hybridization processing of the arrays, or as result of increased steric hinderence conferred by the close proximity of the mismatch to the platform surface. Mismatch probes are useful to determine which miRNAs are especially prone to cross-hybridization on a given array platform, and therefore require further validation by a different method, such qRT-PCR.
5.3.3
MiRNA Preparation from CNS Tissues for Microarray Analysis
Central to all microarray experiments is the ability to isolate good quality RNA and microarray analysis of miRNAs is no exception. Many RNA isolation methods exist but they have all been optimized for capturing longer transcripts (mRNA) as short RNA species were considered unimportant. Since the discovery of miRNAs and other biologically relevant small RNA species, such as siRNAs, piwiRNAs and rasiRNA, conventional isolation methods have been re-designed or new methods of isolation implemented.
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Most early studies relied on total RNA extraction in Trizol to yield miRNAs for microarray analysis [4, 35, 45]. However, due to issues arising from non-specific interaction as a result of hybridization to pre-miRNAs, homologuos regions on target mRNA, and other RNA species a size fractionation step is generally employed to isolate or enrich for small RNA species. In early studies fractionation was carried out using 12–15% denaturing-PAGE [8, 48, 60, 68] or by filtration through size restriction columns such as the Amicon YM-100 (Millipore, Bedford, MA, USA) [5]. However, the need to isolate large numbers of highly reproducible samples for array analysis has driven the development and marketing of several commercial kits and instruments designed specifically for miRNA isolation. One of the most widely used commercial kits for isolating miRNAs is the mirVana™ miRNA isolation kit (Ambion, Autin, Texas, USA) that employs a combination of both chemical extraction and solid-phase extraction [62]. Specifically, organic extraction is followed by immobilization of RNA on glass-fiber filters to purify either total RNA or small RNA species (≤200 nt). Organic extraction involves disruption of the sample in a denaturing lysis buffer followed by acid-phenol:chloroform extraction of the total RNA from other contaminating bio-molecules including DNA. Solid-phase extraction relies on salt and alcohol to decrease the affinity of the RNA for water and increases its affinity for the solid support used (glass-fiber). Small RNA species are then enriched from total RNA by two sequential additions of 100% ethanol followed each time by passage through the solid support. Upon the addition of ethanol the concentration of the alcohol in the preparations increases from 25% to 55%. In the final step, the RNA is washed several times and eluted in low ionic strength solution. MiRNA isolation from CNS tissues by the mirVana kit for microarray analysis has been used in several studies [28, 57, 62]. Some commercial vendors offering miRNA isolation/enrichment kits include Invitrogen’s PureLink™ miRNA isolation kit from (Carlsbad, CA, USA), Qiagen’s miRNeasy Mini Kit (Valencia, CA, USA), Kreatech Diagnostics miRacULS II miRNA isolation and labelling kit (Amsterdam, Netherlands). Denaturing-polyacrylamide gel electrophoresis is a robust, although timeconsuming, methodology for the isolation of small RNAs. Ambion has automated this methodology and introduced the flashPAGE fractionator system that can rapidly and reproducibly fractionate total RNA. Gel fractionation enriches the RNA population ranging between ∼15–40 nucleotides in length by ~10,000-fold. The advantage of this over chemical/solid phase-extraction is that mature miRNAs can be specifically isolated whereas the chemical/solid phase-extraction extraction results in the enrichment of many small RNAs under ~200 nt, not just mature miRNAs.
5.3.4
Labelling miRNAs for Detection via Microarrays
Due to the small size of miRNAs, the lack of common sequence feature(s) and the relatively low amounts present in biological samples, specialized labelling methods are required to achieve consistent and representative labeling of miRNA targets. Most of the methodologies in use have been developed for the analysis of larger transcripts
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such as mRNA and adapted for use with small RNAs. The methodologies currently widely used for labeling miRNAs for array analysis can be classified into two major categories. Either miRNAs are isolated and labeled directly, or an intermediate step, for example reverse transcription or amplification, is employed and results in indirect labeling of the target. Following indirect labeling methodologies the resulting labeled species may be in either the sense or the antisense orientation. Additionally, a number of novel labeling techniques have been developed recently to specifically address the challenges of working with very small and rare targets; these will be described also.
5.3.4.1
Direct Labelling of miRNAs
Several different types of direct labelling strategies have been reported in studies to profile miRNAs from CNS tissues. These strategies either involve the direct labelling of nucleotides within the mature miRNA, or the addition of a labelled-tag to either the 3′- or 5′-terminus of the miRNA. Labelling of the 5′-end of miRNAs with the sensitive radioisotope 32P (γ 33P dATP) using T4 polynucleotide kinase has been used in a number of experiments [35, 67]. In these cases, the labeled-miRNAs were hybridized to nylon membrane arrays, and although relatively large amounts of RNA were used as starting material (5–50 µg), the method allowed the detection of
