Dealing with Interferon Response When Doing RNAi

Off-target effects are a major problem when using RNA interference (RNAi) to silence genes in mammalian systems. One potential source of off-target effects, by either transfected siRNA duplexes or transcriptionally expressed shRNAs, is the inadvertent activation of the interferon response. There are several steps that can be taken to deal with this problem.

Delivery
Interferon response is more likely when high levels of siRNA are used; it is important to transfect the minimum amount of the siRNA duplex that gives rise to a specific RNAi response, as assessed by the level of expression of the target mRNA and/or protein. The level of stable shRNA expression achieved by using lentiviral or retroviral vectors is comparatively modest. Unless very high levels of shRNA expression are achieved, for example, by using highly transfectable cells and a very efficient shRNA expression plasmid, nonspecific activation of the innate immune response are less likely to be induced.

Design
Previous work has shown that the interferon response is induced by dsRNAs of ?30 bp in length and that perfect dsRNAs of as little as 11 bp in length can produce a weak induction. One possible approach to solving the problem of nonspecific activation of the cellular interferon response is to design the siRNA duplex or shRNA precursor so that it does not contain any stretches of perfect dsRNA of ?11 bp.

Detection
If activation of the interferon response remains a concern, it is possible to routinely check for this effect during the course of an RNAi experiment. Analyzing the level of expression of an interferon-response gene, such as oligoadenylate synthase-1 (OAS1), interferon-stimulated gene-54 (ISG54), and guanylate-binding protein (GBP), in the transfected or transduced cells by northern blot or RT- PCR assays are commonly used.

Can there be any more convenient alternative method for checking interferon response? One potentially useful product could be HiTiter™ pre-packaged lentiviruses that would have a fluorescent protein (mTFP1, mWasabi, or the brightest FP in lanYFP) under the control of an ISRE (IFN-stimulated response element) or GAS (IFN gamma-activating sequence)*. This could be another group of Product-on-Demand type of reagents, meaning that we will have the design ready, but only to produce them upon ordering. This way the cost to us and the price to customers can be kept at minimum.

*The expression of the interferon-stimulated genes (ISGs) is induced by the type I interferons IFN-alpha and IFN-beta. A cis-acting element (TAGTTTCACTTTCCC, nucleotides -101 to -87) has been identified in its promoter of one of these genes, ISG54. This element is responsible for the inducible expression of the ISG54 gene and is referred to as IFN-stimulated response element. The human guanylate-binding (GBP) gene is induced by INF-gamma in fibroblasts within 15 minutes of treatment. An IFN gamma-activating sequence (GAS) has been identified in the GBP promoter (nucleotides -123 to -103). To create the interferon reporters, we would insert five direct repeats of this ISRE and/or four direct repeats of this GAS upstream of the basic promoter element (TATA box) and mWasabi GFP gene of the Allele’s patented pLico lentiviral plasmid backbone.

It should be noted, however, that simple transfection of cells with expression plasmids can induce low-level activation of the interferon response, presumably owing to the presence of cryptic convergent promoters that cause the expression of low levels of dsRNA. In general, very low-level activation of the interferon response, that is, activation that exerts a global inhibitory effect on protein translation of less than twofold, is unlikely to be a problem as long as the specificity of any observed phenotype is fully confirmed.

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Tags: dsRNA, expressed shRNA, interferon, miRNA, off-target, RNA interference, RNAi, RNAi toxicity, siRNA

RNAi Therapy Mediated by Linear DNA Cassettes

RNA interference (RNAi) has been demonstrated to be a powerful tool to silence gene expression. Therapies based on RNAi are being developed in numerous application areas at fast paces. Although in basic research both expressed and synthetic double-stranded RNA molecules are broadly used to induce gene silencing, synthetic small interfering RNAs (siRNAs) are deemed easier to deliver in preclinical and clinical studies. Compared to synthetic siRNAs, DNA cassettes that express small hairpin RNA (shRNA), microRNA (miRNA), or strands of siRNAs have advantages of prolonged effects.

RNAi-expressing DNA cassettes have been incorporated into viral and non-viral vectors for delivery. Viral vectors for RNAi carry the same risks as those for gene therapies, and are currently not the method of choice for human therapies. Non-viral DNA molecules, often in the form of plasmids, can be easily created and reproduced, but their efficacy is hindered by delivery barriers at the tissue, cell, and the nucleus levels. These difficulties are in part due to the plasmids’ large size, presence of antibiotic resistance genes, and immuresponse-generating CpG islands created in bacteria during propagation.

One way to alleviate these difficulties with non-viral DNA vectors for RNAi is to use linear DNA cassettes. Linear DNAs traverse nucleopores efficiently. The DNA molecules can be conveniently produced by PCR reactions without going through production in bacteria, avoiding DNA modifications such as CpG motifs and the need for replication origin or drug-resistance genes. Linear DNA encompassing a promoter, coding region, and poly(A) signals has been used for protein production. Similarly, by incorporating a miRNA cassette into linear transcription unit driven by a Pol II promoter was used to express RNAi for inhibiting HBV (Chattopadhyay et al. (2009). There are now available technologies and commercial services (e.g. Vandalia Research, Inc.) to produce therapeutic grade linear DNA by specialized PCR reactions.

Allele Biotech’s patents on DNA-expressed RNAi provide a platform for highly express shRNA or siRNA from a DNA molecule as short as fewer than 200 basepairs, potentially more suitable for large scale production, and even more efficient transduction trough tissue, cell membrane, and nuclear pores than the large linear cassettes used by Chattopadhyay et al. A set of experiments similar to the cited HBV studies could quickly lead to the validation of a possibly the most effect way yet for RNAi therapeutics.

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Tags: Allele RNAi, gene silencing, miRNA, non viral vector, PCR DNA manufacture, pol II promoter, pol III promtoer, RNA interference, RNAi patent, RNAi therapeutics, RNAi therapy, shRNA, siRNA, viral vector

3 Ways of Making DNA Libraries through Oligo Synthesis

Pools of DNA molecules of related but non-identical sequences are often used for selecting cDNAs that encode polypeptides with desired functions (such as in antibody screening), or DNA segments as protein binding sites (through SELEX), or DNA molecules that can catalyze reactions (DNA enzymes or deoxyribozymes), etc. The most direct way of creating DNA libraries is to introduce mixed bases during the synthesis of the oligos that will be used in creating the libraries.

1) The most commonly used method of generating degenerate oligos is to use mixed phosphoramidites (aka amidites, the building blocks of oligo synthesis) at desired positions in an oligo, e.g. using “N” to incorporate dA, dC, dG, and dT nucleotides, or “Y” for pyrimidines, “R” for purines. Mixed base oligos from most oligo suppliers are simple to order (and at no extra charge from Allele and a few other sources). During automated chemical synthesis of oligos, the synthesizer consecutively adds dT, dA, dC, or dG in the case of “N” at a pre-set ratio (e.g.25% each). This procedure does not always result in expected usage of each amidite because different amidites have different coupling efficiency, and the order of addition may also bias against amidites that are added later.

2) Using mixed bases like in method 1) leaves little control to achieve ratios of codons for specific amino acids. On the other hand, by using trimer amidites, which can be used for adding 3 nucleotides in each synthesis cycle, one can create oligos encoding selected amino acids at pre-determined percentages. However, this procedure is difficult to perform because trimer amidites are bulky and hard to couple to the elongating oligo; any moisture present during synthesis would have even more severe adverse effects than with regular amidites. Trimer oligo synthesis projects cost several thousand dollars per oligo on materials alone, and the risk is quite high that the oligos would not turn out of desired properties and qualities. For commercial users, this process has another problem—it is patented.

3) Another method for making library oligos is the so called “split-and-pool”, which is particularly suitable for having diversified amino acids embedded in otherwise common sequences like the CDRs within antibody variable regions. The latest oligo we made last month was a ~72 nt oligo with 8 locations that have pre-determined composition of amino acids, i.e. 20% Ala, 10% Gly, 12% His, etc. The procedure took us about 8 hours and we estimated the cost to be about $1,000. The subsequent sequencing results confirmed that ~70% of the clones using this oligo have desired degeneracy, compared to a similar oligo made by a bigger oligo company, at only 40%. In addition, we did not see any stop codon interruptions or major abnormalities.

DNA pools can also be generated by error-prone PCR, or more specifically with overlapping PCR using degenerate primers. The bottleneck for a library screening is how to handle big enough a number of colonies to accommodate the population, e.g. 10e10, or at least 10e8 clones are needed for finding high affinity antibodies. The second critical point is to have a robust and consistent selection readout such as fluorescence in cell sorting.

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Tags: antibody, CDR, degenerate oligos, DNA library, DNA primer, fluorescence, mixed bases, oligo, oligos, split-and-pool, trimer oligo

How do you produce your iPS cells?

From AlleleForum: First off, thank you for choosing Allele Biotech for your iPSC experiment needs. Now onto your questions

You asked Q1: How many human fibroblast cells you normally to start for transfection. I understand you use 12-well plate? How many days you wait till the cells grow confluent? If the cells never grow confluent, should I still transfer them to feeder plate? Is it critical for the cells to reach confluent, if it is, could you suggest the reasons to me

We usually plate at 70% or about 10e4-10e5 cells and transduce the cells for 2-3 days. It should become confluent in 2-3 days. There is no need for the cells to become confluent before splitting onto feeder cells. Please note for primary cells, do not wait for the cells to get too confluent because contact inhibition may induce growth senescence before cells are reprogrammed.

Q2: How many cells you plate on the feeder plate, let’s say it is 6-well plate, and how many clones would normally pup out from each well?

From one well of a 12 well plate, you can plate 1/5 onto a well of a 6 well feeder cell plate. From there, you should get plenty of colonies.

Q3: At the time when you need to cut the Loxp sites, what passage number you do, do you have to dispense the iPS into single cell? Do you have a detailed protocol for that? Other than virus, do you have any other means to do the job, like plasmid?

Never dispense iPSC into single cells. They do not grow back well if split into single cells. iPSC colonies should be passaged in patches of cells. To excise loxP, the suggested timing is after 12-14 days when the cells are reprogrammed into iPSC colonies. Just transduce the iPSC colonies with Cre virus.

Q4: Is it true, that the 4-in-1 is more powerful than individual ones? Do you have the construct(4-in-one) for sale?

The 4-in-1 is somewhat more effective than 4 individual ones. For license issues, we do not distribute the construct to customers because we only offer packaging service. Similar type of plasmid DNAs may be accessible from other sources.

If you have any other questions or concerns, please let us know. Thanks again.

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Tags: 4-in-1, feeder cells, iPS, iPS colonies, iPS medium, iPS protocol, lentivirus, loxP Cre, splitting iPS cells

From iPSC to induced beta-cells, iN and iCM: dedifferentiation vs direct reprogramming

The success of inducing pluripotency in primary fibroblasts and other cells with a combination of only a small number of transcription factors suggested that fully differentiated cells might change fate following similar treatments. Since the demonstration of induced pluripotent stem cells (iPSCs), at least three examples have been published where 3 cell type-specific factors were selected from a pool of 10-20 candidates that, when expressed from viral vectors, could induce beta-cells, neurons, or cardiomyocytes.

Induced beta-cells [1]: Ngn3, Pdx1, and Mafa, adenovirus injected to in vivo targets

Induced neurons (iN) [2]: Ascl1, Brn2, and Myt1l, lentivirus infecting mouse embryonic fibroblasts (MEF) or tail tip fibroblasts (TTF)

Induced cardiomyocytes (iCM) [3]: Gata4, Mef2c, and Tbx5, lentivirus infecting cardiac fibroblasts or TTF

In all 3 cases, the change of fate seemed to be via direct conversion, without passing through a progenitor cell fate before further differentiation. Like iPSC reprogramming, direct reprogramming also requires a transient supply of inducing factors. Unlike generating iPSCs, the percentage of cells getting reprogrammed is much higher in direct reprogramming, ~20% in the cases of iN and iCM vs 0.1-1% in iPSC. It is likely that a transient, inductive expression of essential factors jump-starts endogenous factors to establish cell fate specific programs; it has also been illustrated that chromatin remodeling through DNA methylation, histone modifications, etc. accompanies the direct reprogramming events.

The requirement of the full complement of inducting factors may vary depending on how close the original cell type is to the new cell type. iPSCs are typically created by using 4 genes, but can be created with just Sox2, Oct3/4 particularly when the cells to be reprogrammed are less differentiated, such as tissue progenitor cells. Instead of a more “complete” direct reprogramming from unrelated cells to iN and iCM, the induced beta-cells come from exocrine cells, which share parental cells with beta-cells.

Looking into the near future, it should be expected that cell type-specific gene expression profiles are being re-examined or created right this moment to look for candidate gene pools specific to other cell types, starting from those with cell therapy relevance. Lentivirus, retrovirus, adenovirus, or baculovirus for mammalian expression are being constructed to carry them into fibroblasts or cells that are close to the end product of direct reprogramming. In a few months, many of these inducing gene-expressing viruses will become shelf products as high titer viruses from suppliers like Allele Biotech, incorporating tools in viral packaging, fluorescent proteins, and polycistronic gene expression systems.

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1. Zhou, Q., J. Brown, A. Kanarek, J. Rajagopal, and D.A. Melton, In vivo reprogramming of adult pancreatic exocrine cells to beta-cells. Nature, 2008. 455(7213): p. 627-32.
2. Vierbuchen, T., A. Ostermeier, Z.P. Pang, Y. Kokubu, T.C. Sudhof, and M. Wernig, Direct conversion of fibroblasts to functional neurons by defined factors. Nature. 463(7284): p. 1035-41.
3. Ieda, M., J.D. Fu, P. Delgado-Olguin, V. Vedantham, Y. Hayashi, B.G. Bruneau, and D. Srivastava, Direct reprogramming of fibroblasts into functional cardiomyocytes by defined factors. Cell. 142(3): p. 375-86.

Tags: direct reprogramming factors, iCM, iN, induced beta-cells, induced cardiomyocytes, induced neurons, iPS, iPS lentivirus, iPSCs, MEF, TTF