A guide to large-scale RNA sample preparation
Analytical and Bioanalytical Chemistry
A guide to large-scale RNA sample preparation
Lorenzo Baronti 0 1
Hampus Karlsson 0 1
Maja Marušič 0 1
Katja Petzold 0 1
0 Department of Medical Biochemistry and Biophysics, Karolinska Institutet , Scheeles Väg 2, 17177 Stockholm , Sweden
1 Katja Petzold
RNA is becoming more important as an increasing number of functions, both regulatory and enzymatic, are being discovered on a daily basis. As the RNA boom has just begun, most techniques are still in development and changes occur frequently. To understand RNA functions, revealing the structure of RNA is of utmost importance, which requires sample preparation. We review the latest methods to produce and purify a variation of RNA molecules for different purposes with the main focus on structural biology and biophysics. We present a guide aimed at identifying the most suitable method for your RNA and your biological question and highlighting the advantages of different methods.
RNA; Sample preparation; In vitro transcription; Chemical synthesis; Preparative high-performance liquid chromatography; Structural biology
In an ever-growing world of new classes of RNAs, the need to
reveal their function and structure is expanding [
]. This need
coincides with the advancement in structural biology methods,
such as the resolution revolution in cryo-electron microscopy
] or the discovery of invisible RNA states by
nuclear magnetic resonance (NMR) methods [
techniques are now available to probe the features of a given RNA,
and each one uniquely accesses structural information at different
resolution, depending on the question posed. The required
sample preparation on a large scale often constitutes the limiting step,
as each structural method strongly relies on the quality (and often
quantity) of the RNA sample that has to be provided (Fig. 1).
Hence first we will give a short overview of the available
techniques to determine RNA structure and dynamics.
X-ray crystallography produces high-resolution data but
relies on RNA samples that are rigid enough to crystallize. It
is often not limited by the size of the sample (Fig. 1) and is
Published in the topical collection Euroanalysis XIX with guest editors
Charlotta Turner and Jonas Bergquist.
therefore commonly used to characterize large protein–RNA
complexes. Cryo-EM has recently undergone a leap in
enhancement of resolution that now rivals X-ray
crystallography, especially for large and rigid molecules (Fig. 1).
Whereas X-ray crystallography relies on crystals analyzed
with high-energy synchrotron radiation, in cryo-EM,
molecules are usually frozen on a surface and single molecules
are observed with a high-resolution electron microscope.
Both X-ray crystallography and cryo-EM have in recent years
delivered detailed insights into ribosome structures [
Fluorescence/Förster resonance energy transfer (FRET) is
based on two fluorophore tags, which can transfer energy on
the basis of their relative proximity. The distance between the
tags can be assessed depending on the efficiency of the FRET
exchange, and this information can then be used to calculate
lowresolution structures. FRET structure calculation requires the tags
be chemically bonded to the molecule, potentially inhibiting
relevant interactions. Most often, more than one donor–acceptor
pair is necessary, requiring several samples. However, this is a
relatively fast method and usually requires a small amount of
sample. FRET is most often used on ribosomes, but will likely
have large use in the localization of regulatory RNA on genomic
DNA or other RNA interactions [
Electron paramagnetic resonance (EPR) can be used to
measure crude long-distance interactions on the basis of a spin
label covalently bonded to the RNA of interest. The method is
in its infancy but can provide information for challenging
systems, where other structural methods fail. In analogy to
FRET, it requires positioning of artificial spin labels that could
potentially interfere with the native structure [
NMR methods provide atomic resolution information,
allow secondary and tertiary structure determination, and make
possible the characterization of molecular motion on a wide
range of timescales [
]. A broad range of sample
conditions can be studied, from dilute solutions to in cell; however,
RNAs larger than approximately 100 kDa are hardly detected
by solution NMR methods . 15N and 13C isotopic labeling
of the sample is a prerequisite when high-resolution structural
data need to be inferred from NMR. With increasing size,
partial or total deuteration of the sample is required.
Small-angle X-ray scattering/neutron scattering measures the
diffraction properties of the atoms in the sample and retrieves
information on the envelope of the molecule. Purified samples
can be measured in native solutions. Small-angle X-ray
scattering/neutron scattering is often used as a complementary
source of information for integrated structural biology studies as
it can easily access the relative position of the components of
large multimolecular complexes [
]. For small-angle neutron
scattering, deuteration of the sample might be necessary.
Secondary structure chemical probing using enzymes or
metal ions has been used for decades, but has recently seen a
revival with small molecules such as selective 2′-hydroxyl
acylation analyzed by primer extension (SHAPE) probing to
reveal the RNA structure [
]. Structure determination by
chemical probing is rather crude, and de novo calculation is
still an exception. Information on secondary structure with
nucleotide resolution is accessible when chemical probing is
performed on purified or native samples in vitro or in vivo,
respectively. It is applicable to large RNAs and protein–RNA
gradients (Table 1). References for relevant examples are indicated to
the right of each structural biology method. Abbreviations: biophys.
biophysical, cryo-EM cryo-electron microscopy, EPR electron
paramagnetic resonance, FRET fluorescence/Förster resonance energy transfer,
NMR nuclear magnetic resonance, nt nucleotide, PCT polymerase chain
transcription, PLOR position-selective labeling of RNA, SANS
smallangle neutron scattering, SAXS small-angle X-ray scattering, sec.
complexes, and can be performed in a high-throughput
fashion. Most protocols still require extraction and processing of
the RNA of interest, and the experimental readout is based on
sequencing or capillary electrophoresis, which can introduce
their own biases.
Biophysical methods, such as UV spectroscopy, circular
dichroism spectroscopy, and isothermal titration calorimetry,
are routinely used as first characterization steps in structural
biology approaches. They are normally performed fast and
require a low amount of purified sample [
], however, they
usually give only a single average signal over the whole
sample of conformations.
Computational prediction is a valuable tool in RNA
structural biology, and the field is fast growing with ever-improved
software. To be able to validate the advancements in the field,
the RNA structure prediction community have created the RNA
puzzle, where different methods compete with and validate
each other every 2 years, providing a great display of which
software is available and how the different types perform [
All of these experimental methods need copious amounts
of highly pure RNA to answer the different questions about
RNA biology and structure. Unfortunately, working with
RNA is hindered by one major hurdle, the ubiquitous presence
of RNases, proteins that degrade RNA at a high rate. This
requires careful and extensive preparation of laboratory
equipment in a so-called RNase-free fashion [
] and/or the use of
different, commercially available, RNase inhibitors.
This review aims to reach out to the inexperienced
reader in the field of RNA structural biology and to
provide a starting point for the sample preparation literature,
including the latest innovations.
Recombinant overexpression in Escherichia coli has
revolutionized protein structural biology by allowing the production of
large quantities of sample from cheap biomass fermentation
and quick recovery from bacterial lysates with use of fusion tags
for affinity purification. In contrast, the development of RNA
heterologous expression has been heavily hampered by a few
crucial factors; namely, the degradation by intracellular RNases,
the large 3′-end and 5′-end heterogeneity of the transcripts, and
the lack of efficient tags for affinity purification [
]. See for
overview Table 1.
Transfer RNA scaffolds
To circumvent such limitations, Ponchon and Dardel [
developed a method that exploits transfer RNA (tRNA) as
a protective scaffold to accommodate target sequences in the
anticodon stem. A similar, although less popular, variant that
uses 5S ribosomal RNA (rRNA) as a scaffold was also
]. In the Ponchon and Dardel method, tRNA
(human tRNALys3 or E. coli tRNAMet) chimeras are cloned
into an expression vector, where the insertion of choice
replaces the anticodon stem, while maintaining the native fold
of the TΨC and D stem-loops. In this way the maturation
machinery processes and correctly folds the chimeras to
homogeneous species that stably accumulate in the host in up
to 50 mg of RNA per liter of culture [
]. The scaffold
can then be cleaved, releasing the insert via a co-transcribed
ribozyme, a DNAzyme, or by annealing of the construct with
a pair of complementary DNA oligonucleotides and
subsequent digestion of the undesired DNA/RNA hybrid with
RNase H . This technique has proven successful for a
broad range of RNA sizes and structures, including the
expression of fusion aptamers for affinity purification [
General approaches to in vivo RNA expression have been
extensively reviewed , including applications to
] and production of small interfering RNA/microRNA
]. A recent review focused on in vivo
sample preparation for high-resolution structural studies
presented the most efficient strategies to design, process, and
separate scaffold RNA from the insert of interest .
Recombinant expression has also been applied to NMR
studies of large RNAs (more than 40 nucleotides) where
expensive atom-specific isotopically labeled nucleotides (2H, 13C,
and 15N) are normally demanded for unambiguous data
11, 47, 48
]. Recently, the Dayie group [
showed that by combination of the tRNA scaffold with two
metabolically different, complementary E. coli strains, it is
possible to achieve cost-effective preparation of 13C
selectively labeled NMR samples in vivo.
A new approach to increase sample stability from exonuclease
degradation was proposed on the basis of circular RNAs, a
novel class of RNAs characterized by the covalent 3′–5′
linkage found in many organisms. The pathways involved in
generation of circular RNAs as well as in vitro methods for
circularization were recently reviewed, including a potential
Advantages and disadvantages of different RNA production methods to aid, in combination with Fig. 1, in the selection of the most suitable
nt nucleotides, PCT polymerase chain transcription, PLOR position-selective labeling of RNA, tRNA transfer RNA
heterologous RNA production [
]. Intronic circular RNAs
generated during tRNA biogenesis in metazoans were also
shown to be effectively used for recombinant RNA expression
in vivo [
Alternative hosts for in vivo production
The possibility to use different expression hosts alternatively
to E. coli is currently being explored (e.g., use of Rhodovulum
sulfidophilum to produce recombinant human precursor
miR29b that is directly retrievable from the extracellular medium)
]. In a recent study, Pereira et al.  compared the
performances of E. coli and R. sulfidophilum, showing that
even though purification from intracellular fractions of
E. coli produces a higher yield with shorter fermentation
times, use of R. sulfidophilum as an expression host drastically
simplifies downstream purification and limits protein
contamination. Although far from being as broadly applicable as the
tRNA scaffold in the E. coli method, the R. sulfidophilum
approach highlights the potential behind noncanonical
expression hosts for future in vivo RNA production.
RNA transcription is one of the three fundamental reactions
that define the central dogma of biology, and the enzymes that
perform such a reaction are ubiquitously expressed among all
life forms. In most cases, RNA polymerases (RNAPs) have
evolved to participate in multimolecular machineries that
require different protein cofactors for each stage of RNA
synthesis (initiation, elongation, and termination). Although
possible, recapitulating transcription in vitro for most bacterial,
archaeal, or eukaryotic systems is still challenging, and the
yields are not suitable for biotechnological applications. In
contrast, bacteriophages have evolved their RNAPs to
optimize genome compactness and maximize transcription yields
during the lytic phase of their life cycle. This results, in the
case of bacteriophages T3, T7, and SP6, in RNAPs that
comprise a single polypeptide chain and require only Mg2+ as a
cofactor to perform highly processive RNA synthesis. These
first observations [
] allowed the development of T7 in vitro
transcription, now the most established method for enzymatic
RNA preparation (Fig. 1).
In brief, T7 RNAP accepts ribonucleotide triphosphates as a
substrate to synthesize a transcript RNA complementary to a
DNA template of choice. Once the enzyme has completed
the polymerization, it runs off the DNA template and
releases the transcript, thereby ensuring the process is
performed several times (Fig. 2a). In this way, milligram
quantities of RNA can be produced per milliliter of transcription
reaction in a few hours. Historically, two major drawbacks
of the T7 system are promoter specificity and heterogeneous
ends. Firstly, RNAP has a strong specificity for the promoter
sequence, which extends beyond the initiation site, and
therefore this limits the possibility to incorporate any
userdefined 5′ sequence (Fig. 2a). Secondly, the inconsistent
runoff of the RNAP leads to 3′-end inhomogeneity of the
transcript, most commonly with a single A extension. To
overcome such limitations, many improvements to T7
in vitro transcription have been made, and excellent
overviews of transcription optimization techniques, including
a T7 in vitro transcription
3’ T7 promoter class III
CH3 R CH3 R’
3’O OO OO O
DN5A’- mtmeemotdhpioflaixetyde O
Position-selective Labelling Of RNA (PLOR)
Polymerase Chain Transcription (PCT)
5’ lSaebleelclitnivge N cycles
3. Termination Transcript
n+m (i.e. 25+18 nt)
Cycle 1 chimeras
94 °C, 15-30 s - Melting
35-49 °C, 1 min - Annealing
50 °C, 1h - Transcription
Fig. 2 Transcription-based sample preparation methods. a T7 in vitro
transcription scheme; the double-stranded (dsDNA) T7 promoter
sequence and 5′-methoxy modification of the DNA template strand are
highlighted. b Position-selective labeling of RNA in the solid and
solution phase (left). Fundamental steps in one cycle that correspond the
production of a single RNA transcript from one DNA template (right).
c Polymerase chain transcription components (DNA template,
asymmetric DNA primers, and SFM4-3 Pol) and reaction cycles. The
RNA is exponentially amplified through thermal cycling transcription
reactions; the DNA–RNA chimeras are subsequently digested to obtain
the final RNA products. Abbreviations: biot biotin, nt nucleotide, NTP
nucleotide triphosphate, RNAP RNA polymerase, SA streptavidin,
ssDNA single-stranded DNA
step-by-step protocols, are available [
]. Recently, the
combination of two previously known, yet unrelated,
approaches to in vitro transcription was shown to greatly
increase 3′-end homogeneity: the use of
C2′-methoxy-modified DNA templates in the last two positions at the 5′ end
of the DNA template reduces incorporation of 3′-end
nontemplated nucleotides  (Fig. 2a), whereas low
concentrations of dimethyl sulfoxide added in the transcription
reaction increase T7 RNAP synthesis yield through an
unknown mechanism [
]. Recent observations showed that
C2′-methoxy-modified DNA templates in combination with
20% dimethyl sulfoxide reduce the addition of nontemplated
nucleotides at the 3′ end to the extent that subsequent
purification from side products was no longer necessary for the
structural studies proposed in the study [
]. This approach
was successfully applied to the transcription of the
wellestablished 5′ leader of HIV genome RNA construct, where
3′-end inhomogeneity impairs the dimerization properties of
the construct, thereby improving sample preparation and
allowing a high-resolution NMR study of the whole
intermolecular interaction site [
Ribozyme cleavage and T4 ligation
Complementary approaches to obtain 5′-end and 3′-end
homogeneous RNAs use the nuclease activity of different
enzymes to trim the transcript to the desired length. Cleavage
of the phosphodiester backbone at a specific site is
commonly achieved by design of a fusion transcript carrying a
cisacting, self-cleaving ribozyme [
]. Most cis-acting
ribozymes co-transcriptionally fold and cleave the backbone
at a specific site with high efficiency and few or no sequence
requirements. When expensive, labeled ribonucleotide
triphosphates are used during transcription, trans-acting
ribozymes can also be used after purification of the RNA
of interest. Hammerhead, hepatitis delta virus, and Varkud
satellite ribozymes are commonly used ribozymes, and their
application, in particular to segmental labeling, was reported
recently . Despite their wide use, one major limit of
hammerhead and hepatitis delta virus ribozymes is the
generation of 5′-hydroxyl and 2′-3′-cyclic phosphate ends,
which requires tedious additional sample treatment when
native 5′-phosphate and 3′-hydroxyl ends are needed [
DNAzymes can also be used as trans-acting ribozymes;
these, however, also generate noncanonical ends and require
a purine/pyrimidine site for cleavage. Alternatively,
canonical 5′-phosphate and 3′-hydroxyl ends can be obtained by
RNase H cleavage after annealing of the target cleavage site
with C2′-methoxy RNA/DNA chimera oligonucleotides.
Recently, the development of new bioinformatic tools led
to the discovery of a new class of ribozymes called
Btwisters^ and comprising twister, twister sister, pistol, and
]. Structural characterization of twisters has
begun to emerge, suggesting interesting potential for the
future . However, a general consensus on the mechanism
of action of these new ribozymes is still lacking [
biotechnological application seems far away at the moment.
5′-Hydroxyl and 2′-3′-cyclic phosphate ends generated by
ribozymes are a useful tool to avoid self-ligation and achieve
the correct order of ligated fragments in segmental labeling
]. Combination of fragments with different
labeling properties is a fundamental approach in the study of
large RNAs (more than 40 nucleotides) by NMR
experiments or for selective placement of paramagnetic and
fluorescent probes for EPR and FRET experiments, respectively.
Most segmental labeling techniques are based on sequential
steps of enzymatic cleavage and ligation of in vitro
transcribed or chemically synthesized RNA oligonucleotides.
Ligation occurs between an acceptor fragment bearing a
3′hydroxyl and a donor fragment with a 5′-monophosphate
end. The ligation reaction can be performed with the T4
DNA ligase with the aid of a DNA splint complementary
to the site of ligation (splint ligation) or by the T4 RNA
ligase, which ligates donor and acceptor ends, which are
brought close together by base pairing (nonsplinted ligation).
A comprehensive review of available methods and detailed
protocols of the most recent segmental labeling strategies
proposed by the Allain group are available [
of these techniques are affected by the inefficiency of T4
DNA ligase, and the final yield is normally low even for a
two-fragment reaction. In the ideal scenario, where yield and
downstream purification are optimized, the protocol can take
up to 7 days to be completed [
]. Moreover, when the
fragments to be ligated are about the same size as the
ribozyme used in the cleavage reaction, purification of the
product of interest can be troublesome.
Position-selective labeling of RNA
An alternative to ligation methods for segmental labeling was
recently developed by the Wang group [
], combining T7
in vitro transcription with the solid-phase synthesis approach.
Position-selective labeling of RNA (PLOR) exploits the
ability to pause and restart T7 transcription by supplementing the
reaction mixture with incomplete nucleotide triphosphate
(NTP) mixes in a stepwise fashion (Fig. 2b). In the original
work, the authors synthesized the 71-nucleotide aptamer
domain of an adenine riboswitch (riboA71) with selective
labeling of the stem, linker, or loop regions. In this case the DNA
template is incubated with T7 RNAP and an NTP mix
(unlabeled) lacking CTP that causes stalling of the
transcription at the position where the first CTP would be incorporated.
Next, the DNA–RNAP–transcript tertiary complex bound to
the solid phase is separated from reagents and cofactors in the
liquid phase through extensive washing steps (Fig. 2b). The
reaction is reinitiated by addition of a new NTP mix (labeled
or unlabeled) that allows elongation of the transcript up to a
new stalling site. Addition of incomplete NTP mixes can be
repeated as many times as the nucleotide sequence allows for
correct incorporation and stalling, and depending on the
number of labeling sites wanted. Once the RNAP reaches the last
stalling point, a full NTP mix is added to terminate the
transcription. The transcripts are collected from the liquid phase,
and reinitiation is inhibited by rapid cooling to 4 °C. The
agarose-bound DNA template can be reused several times,
and the whole sequence of initiation, elongation, and
termination can be repeated for N cycles (Fig. 2b). Because of the
many incubation/washing steps involved, a fully automated
platform that allows fast and efficient PLOR synthesis was
]. This approach has proven successful for the
production of different samples of riboA71 for NMR studies,
single molecule FRET studies [
], and X-ray free
electron laser serial crystallography.
Polymerase chain transcription
The iconic production of DNA via polymerase chain reaction
amplifies exponentially the initial templates through thermal
cycling reactions with high efficiency and minimal side
products. Although in principle possible, RNA amplification via
polymerase chain reaction has never been developed, mainly
because of the lack of efficient heat-resistant RNAPs. A
different approach was used by Cozens et al. [
], changing a
DNA polymerase into an RNAP. To allow substrate tolerance
of Thermococcus gorgonarius DNA polymerase, the group
developed a mutant enzyme capable of synthesizing RNA
and modified nucleic acids while retaining its stability toward
heat denaturation [
]. The Romesberg group recently
reported a variant of the Stoffel fragment of the Taq DNA
polymerase (SFM4-3) [
] that was shown to exponentially amplify
two different RNA fragments from one DNA template,
primed by two asymmetric DNA oligonucleotides [
2 c ) . T h e r e a c t i o n w a s n a m e d Bp o l y m e r a s e c h a i n
transcription^ and has the potential to exceed T7 transcription
in terms of amplification levels (102–104-fold reported). It
allows incorporation of 2′-F-modified NTPs and, because of
the increased reaction temperature in comparison with in vitro
transcription, allows a more efficient transcription of
structured DNA templates with high GC content.
Although generally dominated by chemical synthesis, the
development of engineered DNA polymerases for the
synthesis of modified nucleic acids is a growing field, and enzymatic
modification of NTPs and RNA oligonucleotides answers the
need for unnatural oligonucleotides in synthetic biology,
imaging, and therapeutics. A detailed description of the available
modification methods is beyond the scope of this review;
however, we direct the interested reader to a series of reviews
that highlight the latest advancements in polymerase
engineering and enzymatic synthesis of modified RNAs [
Currently, solid-phase chemical synthesis is the method of
choice for the production of oligonucleotides shorter than 10
nucleotides, as these cannot be produced efficiently by the T7
system (Fig. 1) [
]. The upper size limit of the method is
approximately 80 nucleotides. If longer constructs are desired,
shorter chemically synthesized RNAs can be linked together
with the help of T4 DNA or RNA ligase in the splint ligation
]. Synthesis proceeds from the 3′ end to the 5′ end,
and involves four steps (Fig. 3a). First, the 5′-hydroxyl
Fig. 3 Solid-phase synthesis of RNA. a The different steps in the
solidphase synthesis cycle of RNA and the most commonly used. b 2′-OH and c
5′-OH protecting groups (PG). At the end of the production cycle, functional
groups that are not participating in the polymerization reaction are
deprotected. Abbreviations: ACE 2′-bis(2-acetoxyethoxy)methyl, DMT
4,4′-dimethoxytrityl, TBDMS 2′-O-(t-butyldimethylsilyl), TC
2′thiomorpholine-4-carbothioate, TOM 2′-O-[(triisopropylsilyl)oxy]methyl
protecting group of the nucleotide that is bound to the solid
support is removed to allow the 5′-hydroxyl to be attacked by
the activated 3′-hydroxyl group of the incoming nucleoside
phosphoramidite monomer during the second, coupling step.
In the third, capping step, unreacted 5′-hydroxyl groups are
blocked from participating in subsequent reactions to avoid
synthesis of side products. Finally, the unstable phosphite
triester formed during the coupling step is converted to a stable
species by an oxidation step, and the whole cycle can be
repeated to obtain the desired length of the oligonucleotide polymer
(Fig. 3a) [
]. Postsynthetic steps include cleavage from the
solid support, which is either polystyrene or controlled-pore
glass, and deprotection of reactive groups of the nucleobases.
Purification of the whole-length product is in principle more
straightforward than with enzymatic methods, as the most
commonly used 5′-hydroxyl protecting group, 4,4′-dimethoxytrityl,
can be left attached to the 5′-end nucleotide and used for
purification of the whole-length product with reversed-phase
highperformance liquid chromatography (HPLC).
The efficiency of solid-phase RNA synthesis depends on
the type of RNA phoshoramidite monomers used, where
currently four different RNA phosphoramidite monomers are
commercially available. All use a mild base-labile protection
on the amino groups of cytosine, guanine, and adenine
nucleobases as well as on phosphate and the support linkage.
Variations exist, however, with 5′-hydroxyl and 2′-hydroxyl
group protection. 2′-O-(t-Butyldimethylsilyl) [
2′O-[(triisopropylsilyl)oxy]methyl  groups are most
commonly used for hydroxyl protection (Fig. 3b, c), and these
allow the use of modifications that are sensitive to
deprotection conditions. However, the biggest drawback of
these moieties is long coupling times and the ability to
synthesize only shorter (up to 60 nucleotides long)
oligonucleotides. For longer RNAs, more expensive phosphoramidite
monomers with shorter coupling times and greater coupling
efficiencies can be used: 2′-bis(2-acetoxyethoxy)methyl
] and 2′-thiomorpholine-4-carbothioate
] (Fig. 3b, c). A main drawback of
2′-bis(2acetoxyethoxy)methyl RNA monomers is the need to use
fluoride to remove the 5′-hydroxyl protecting group, requiring
modifications of current RNA synthesizers [
]. In contrast to
all other types of phosphoramidite monomers, all
2′thiomorpholine-4-carbothioate protecting groups can be
removed in the same conditions, which makes RNA synthesis
more convenient and less time-consuming. For a more
detailed review of the current reactions used, see [
Solid-phase chemical synthesis of RNA allows the
introduction of isotopically labeled groups or chemical modifications at
specific positions directly during the synthesis or through a
postsynthetic labeling reaction with a reactive group [
This is especially useful for techniques that require site-specific
labeling schemes for simplification of spectra (NMR), need the
incorporation of dyes and florescent probes (FRET), or require
precise positioning of spin labels (EPR) (Fig. 1). The limitation
for incorporation of modified and/or labeled residues in
chemically synthesized RNA lies only in obtaining the appropriate reactive
phosphoramidite monomers. A subset of monomers with
chemically modified or fluorescent groups are commercially available,
and several groups are working on new modified or labeled RNA
building blocks that have been successfully incorporated with
classical solid-phase synthesis [
]. Lately, improvements in
isotope labeling of RNA, especially site-specific deuteration and
segmental labeling of RNA, have opened the avenue for studying
RNA molecules of ever-increasing size particularly by NMR
spectroscopy . For example, a novel strategy for resonance
assignment that combines new strategic 13C labeling technologies
with filter/edit-type nuclear Overhauser effect spectroscopy
experiments to greatly reduce spectral complexity and crowding was
proposed very recently [
]. This new strategy allowed
assignment of important nonexchangeable resonances of proton and
carbon nuclei with only one sample and less than 24 h of NMR
instrument time for a 27-nucleotide-long model RNA. Another
example includes the use of site-specific labeling of the ribose C1′
or C2′ position that allows measurements of dynamics with use of
Carr–Purcell–Meiboom–Gill sequences [
Precipitation and solvent extraction
Selective isolation of RNA from complex mixtures is a step
needed in most purification protocols to achieve a first, rough
refinement of the sample before more sophisticated separation
techniques. Precipitation and solvent extraction methods take
advantage of differential solubility of biomacromolecules in
different solvents and ionic conditions. Precipitation is
generally performed after in vitro enzymatic reactions to separate
the RNA of interest from the protein and DNA components or
simply for buffer exchange, whereas solvent extraction
followed by precipitation is the method of choice to isolate
large amounts of total RNA from natural sources. See for
overview Table 2.
The polar nature of the negatively charged backbone makes
RNA highly soluble in water. Several cations used in
combination with ice-cold ethanol as a co-solvent can effectively
neutralize the backbone charges and reduce the solubility to
a point where the RNA selectively precipitates out of solution.
Different cations and their salts, such as ammonium acetate
and lithium chloride, can be used depending on the size and
concentration of the RNA to be precipitated. A comprehensive
method comprising step-by-step protocols for RNA
precipitation can be found in [
Advantages and disadvantages of different RNA purification methods
AC affinity chromatography, HPLC high-performance liquid chromatography, IE ion exchange, IP ion paring, PAGE polyacrylamide gel electrophoresis,
RP reversed phase, SEC size-exclusion chromatography
RNA isolation by acid guanidinium thiocyanate–phenol–
chloroform extraction was initially developed as an alternative
to the tedious total RNA isolation from mammalian tissues
using ultracentrifugation. In this method the sample is
incubated with an equimolar mix of phenol and chloroform, which
allows proteins to be denatured by the guanidinium
thiocyanate and consequently separated in the organic phase, whereas
RNA is dissolved in the aqueous phase. Separation of the two
phases is then achieved by centrifugation, and RNA can be
retrieved by subsequent ethanol or lithium chloride
precipitation. In acid guanidinium thiocyanate–phenol–chloroform the
polar phase is kept under acidic conditions (pH 4–6), allowing
the RNA to remain soluble while the DNA undergoes
repartition at the interface. A detailed and reedited version of the
original protocol was published [
], and many commercial
kits based on this method are available.
Ultracentrifugation is the standard way of purifying large
RNA machines, such as ribosomes and ribosomal subunits,
as well as other macromolecules of biological origin in the
same size range. Ultracentrifugation with a gradient of a solute
such as sucrose has been used for more than half a century
]. For the isolation of complete ribosomes, polysomes, or
individual ribosomal subunits, large amounts of cells from the
organism of interest are mechanically lysed, the lysate is
subsequently centrifuged at low speed to get rid of cell debris, and
is then ultracentrifuged at high g (approximately 105g) on top
of a sucrose cushion to form a pellet of the required ribosomes
]. The material can then be further purified by
ultracentrifugation on a variety of buffered sucrose gradients with
different salt content. The ability of the ribosome to keep its
two subunits together is highly magnesium dependent, and by
tuning the Mg2+ content of the sucrose gradient used for
purification, one can obtain complete ribosomes or individual
subunits. A lower magnesium ion concentration (1 mM) will aid
in separating the individual subunits, whereas a higher
concentration will promote isolation of complete ribosomes [
After separation of the ribosomal components, the UV
absorbance of the content from the centrifugation tubes is measured,
and the content is fractionated. From the absorbance profiles,
the fraction content can be deduced [
traditionally heavily used within the field of structural biology in X-ray
crystallography and cryo-EM, the ultracentrifugation
technique has also found its way into other fields of research.
Multiple methods where ribosomes are isolated for
investigation of the messenger RNA being translated at a specific
moment, the translatome, also use the ultracentrifugation
]. Ultracentrifugation has also proven useful for
nucleic acid related nanotechnology or DNA origami, where
ultracentrifugation onto glycerol gradients has been used as a
good complement to more conventional and established
agarose gel electrophoresis purification methods .
Polyacrylamide gel electrophoresis
For decades, polyacrylamide gel electrophoresis (PAGE) was
the standard method to purify large amounts of RNA
(microgram to milligram scale) with single-nucleotide resolution as it
can be easily applied to a wide range of RNA sizes and
requires a minimal setup with cost-effective reagents. Like all
electrophoretic techniques, PAGE uses a polymeric mesh to
separate molecules by size, and/or conformation, as charged
macromolecules migrate through an electric field. The mesh is
prepared by polymerization of an acrylamide/bisacrylamide
solution. The concentrations of acrylamide/bisacrylamide
monomers can be varied to obtain the desired pore size and
resolving power in the final mesh. Typical monomer
concentrations range from 5% to 20% for RNA gels with 1:19
acrylamide/bisacrylamide relative ratio. In this range, short
to intermediate length (5–500-nucleotide) RNAs can be
]. The acrylamide solution can be supplemented
with a denaturing agent (most commonly 8 M urea, called
Bdenaturing PAGE^) to fully unfold the migrating RNA and
separate the molecules solely by size. With this approach,
single-nucleotide resolution can be easily achieved in
largescale preparations, and it is a common analytical method.
When no denaturing agent is used and the electrophoresis
apparatus is externally cooled to prevent overheating caused
by electric current flowing through the gel, conformers with
different hydrodynamic radii of a given RNA construct can be
resolved; this is also the underlying principle of
electrophoretic mobility shift assays [
]. Isolation of the RNA of interest
is obtained by excision of the band of interest from the gel,
followed by electroelution or crush and soak extraction [
Gel excision is routinely used, and requires only a UV lamp to
identify the band of interest by UV shadowing on fluorescent
paper and a scalpel to excise the band. The excised band is
then treated to allow diffusion of the RNA from the gel mesh
into solution, and the RNA is subsequently purified by ethanol
precipitation and resuspended in a buffer of choice. The
impact of UV shadowing on RNA integrity has recently been
addressed, revealing that commonly used lamps and exposure
times can lead to minimal but detectable photodamage of the
sample, and thereby potentially corrupt downstream structural
]. Electroelution requires a dedicated apparatus
that allows collection of the species corresponding to the band
of interest being eluted from the gel pieces. RNA preparative
PAGE has been used for decades, and its use has been well
described in textbooks; however, a more comprehensive
stepby-step protocol has recently been published with a focus on
RNA preparation for structural biology application [
Liquid chromatography techniques are well established for the
purification and analysis of nucleic acids and oligonucleotides
spanning a wide size range. On the basis of the concept of
allowing solutes in a solvent (mobile phase) to run through a
column containing a solid material (stationary phase) that
interacts with the solutes to different extent, separation is
]. Here we focus on the chromatographic
methods that are relevant for RNA sample preparation (Fig.
4). Among the sorptive separation techniques commonly used
for nucleic acid separation are reversed-phase ion-pairing
HPLC (RP-IP-HPLC), ion-exchange HPLC (IE-HPLC), and
ion-exchange fast-performance liquid chromatography
]. Lately, affinity chromatography methods,
falling within the category of sorptive techniques, have been
increasingly used for RNA preparation. Further, there are
examples of successful RNA purification approaches using
sizeexclusion chromatography (SEC), where the molecules are
separated on the basis of molecular size rather than chemical
interactions with the stationary phase [
]. All of these
techniques and their relevance for RNA sample preparation have
been reviewed recently [
]. A more detailed
description of the different methods follows.
Fig. 4 Liquid chromatography methods. a Reversed-phase ion-pairing
chromatography; the lipophilic stationary phase retains the RNA thanks
to a lipophilic cation-pairing agent (tetrabutylammonium is depicted). b
Ion-exchange chromatography; a positively charged stationary phase
interacts and retains the negatively charged RNA molecules. c Affinity
chromatography; a polyuridine (poly(U)) functionalized stationary phase
selectively interacts with polyadenosine (poly(A)) tails of messenger
RNAs (mRNA). d Size-exclusion chromatography; large RNAs are
eluted through the porous medium of the stationary phase with short retention
times, and smaller RNAs are absorbed into the porous medium, resulting
in longer retention times
O O R
O O R
3’- A A A A
U U U UU U
Reversed-phase ion-pairing high-performance liquid chromatography
This technique is based on the use of lipophilic cations; quaternary
ammonium compounds that ion-pair with the negatively charged
sugar–phosphate backbone of the oligonucleotide are commonly
used. These ion-paired complexes then become lipophilic and can
interact with the stationary phase of a reversed-phase
chromatography column (Fig. 4a). The lipophilic oligonucleotide complex,
once bound to the column, is eluted and separated with an organic
solvent gradient, usually with acetonitrile [
]. As described
], although RP-IP-HPLC has been used
successfully for oligonucleotide separation for almost 40 years for smaller
amounts of material (analytical scale), scaling up of the method is
rare (preparative scale). Examples of reversed-phase ion-pairing
methods, purifying larger amounts of oligonucleotides, in the
preparative scale of around 1 mg can be found [112, 113], but
for these and similar examples the material purified is exclusively
synthetic oligonucleotides, which are already inherently pure.
RPIP-HPLC methods are commonly developed for analytical
purposes [114, 115]. Some recent work on the analytical scale might
be also applicable to RNA sample preparation: for example,
RPIP-HPLC has been used to analyze and purify double-stranded
RNA (dsRNA) from material expressed in E. coli . In this
work total RNA was extracted from the bacteria and analyzed by
RP-IP-HPLC. Advantage was taken of the fact that
singlestranded RNA can be subsequently degraded to isolate the
dsRNA. This method exemplifies the usability of RP-IP-HPLC
to assess the presence and purity of dsRNA as well as the native
RNA of the cell (e.g., separation of tRNAs and the different
ribosomal RNAs). Another method addresses the separation of
stereoisomers to synthetically produce RNAs containing
phosphorothioate groups in the backbone by RP-IP-HPLC .
This work shows that it is possible to separate stereoisomers using
classic reversed-phase ion-pairing chemicals and ion-pairing
agents. Triethylamine acetate in combination with the organic
modifier acetonitrile was the most successful combination.
Ion-exchange high-performance liquid chromatography
Just like the RP-IP-HPLC method, IE-HPLC and IE-FPLC are
equally established as a technique for oligonucleotide separation.
In this technique the stationary phase already contains the
cationic groups for the anionic oligonucleotide to interact and bind with
(Fig. 4b). The polymer is then eluted and separated on the
column with use of a salt gradient [
]. IE-HPLC has been used
successfully within the field of RNA structural biology, and
] might be one of the most attractive options to
purify milligram amounts of in vitro transcribed RNA. However,
even with this method, abortive transcripts can be present in the
final purified material for molecules shorter than 30 nucleotides
when one starts with a heterogeneous RNA sample. IE-HPLC
has also been successfully used together with trans-acting
hammerhead ribozymes (see BRibozyme cleavage and T4
ligation^) to purify in vitro transcribed RNA on a milligram scale
]. The fact that the ribozyme is trans-acting can make this
approach attractive to purify isotopically labeled RNA samples,
since the cleaving ribozyme can be transcribed unlabeled in a
completely independent transcription reaction. Several important
aspects of IE-HPLC have been reviewed concerning how
different cations support different conformers of RNA that
subsequently influence the IE-HPLC separation .
Affinity chromatography techniques
As stated before, the affinity chromatograph technique falls
within the category of sorptive techniques; that is, the
compound to be separated interacts chemically with the stationary
phase (Fig. 4c). The major difference compared with
ionexchange and reversed-phase chromatography is that the mode
of interaction between the analyte and the stationary phase is
strongly specific and is often inspired by biological interactions
. The type of molecules in the stationary phase, the affinity
ligands, which interact with the molecule to be separated in the
mobile phase, can differ widely: antibodies, proteins,
oligonucleotides, dyes, boronate groups, or chelated metal ions are
attached to a solid support such as agarose beads or silica
material and constitute the stationary phase . This versatility
of the stationary phases has led to a number of different RNA
purification approaches, some of which were reviewed recently
]. Unless the RNA to be purified naturally contains a
sequence with strong affinity for a target that can be immobilized
on the stationary phase, the RNA can be tagged with a specific
sequence to do so, analogous to the polyhistidine tag used in
protein science. There are several ways to achieve this. Affinity
tags can be developed by systematic evolution of ligands by
exponential enrichment (SELEX) . One approach
developed RNA sequences binding streptavidin and Sephadex. The
tagged RNA was eluted from the affinity ligand by competitive
binding of the natural ligand D-biotin or dextran . In
another elegant approach the tag can be combined with
compound-activated ribozyme to cleave the product of interest
from the stationary phase .
As the name implies, SEC separation is based on the
difference in size or hydrodynamic radius of the molecules. When a
porous stationary phase is used, large molecules cannot enter
the pores and they pass through the stationary phase, whereas
smaller molecules are retained (Fig. 4d). On the basis of this
principle, separation is achieved, and the larger molecules are
eluted first [
]. SEC systems have been successfully used
for preparative-scale RNA sample preparation [125, 126].
Important effects of RNA structure, such as the influence of
oligonucleotide secondary structure and its influence on SEC
separations, have been investigated . For
phosphorothioate oligonucleotides, SEC separation properties have recently
been investigated . This work concluded that
phosphorothioate oligonucleotides can be efficiently separated by SEC
although this is complicated by additional lipophilicity caused
by the sulfur atoms in the backbone of the oligonucleotide.
Sample preparation is currently a bottleneck in the structural
characterization of RNA, impeded by RNases and a lack of
development for large-scale preparative methods. However,
new developments are opening up new avenues that can be
combined to answer many new, challenging, and interesting
questions in RNA structural biology. Here we reviewed recent
progress in well-established production and purification methods
that allows preparation of large amounts of RNA commonly
needed for structural characterization. Several new exciting
methods that are emerging, such as circular RNAs, PLOR, and
polymerase chain transcription, have great potential to reduce the
amount of work and time needed for RNA sample production.
On the other side, obtaining a sample that is pure and
homogeneous is usually the most challenging step in structural studies,
and new, fast, and innovative methods for sample preparation are
currently lacking. However, a combination of typically two
different purification methods will most often result in a reliable,
easy to work with sample that will save time in the long run.
Acknowledgements We thank J. Schlagnitweit and E. Johnston for
critical reading. We acknowledge the Karolinska Institutet for PhD student
support for LB (DNR 2-3707/2013). We are grateful for the financial
contributions from Vetenskapsrådet (#2014-4303), Stiftelsen för
Strategisk Forskning (ICA14-0023), Jeansson Stiftelsen (JS2015-0126),
and the Ragnar Söderbergs Stiftelse (M91-14).
Compliance with ethical standards
Conflict of interest The authors declare that they have no competing
Open Access This article is distributed under the terms of the Creative
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distribution, and reproduction in any medium, provided you give
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Creative Commons license, and indicate if changes were made.
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