A Robust Protocol for Efficient Generation, and Genomic Characterization of Insertional Mutants of Chlamydomonas reinhardtii.

Abstract

Background
Random insertional mutagenesis of Chlamydomonas reinhardtii using drug resistance cassettes has contributed to the generation of tens of thousands of transformants in dozens of labs around the world. In many instances these insertional mutants have helped elucidate the genetic basis of various physiological processes in this model organism. Unfortunately, the insertion sites of many interesting mutants are never defined due to experimental difficulties in establishing the location of the inserted cassette in the Chlamydomonas genome. It is fairly common that several months, or even years of work are conducted with no result. Here we describe a robust method to identify the location of the inserted DNA cassette in the Chlamydomonas genome.

Results
Insertional mutants were generated using a DNA cassette that confers paromomycin resistance. This protocol identified the cassette insertion site for greater than 80% of the transformants. In the majority of cases the insertion event was found to be simple, without large deletions of flanking genomic DNA. Multiple insertions were observed in less than 10% of recovered transformants.

Conclusion
The method is quick, relatively inexpensive and does not require any special equipment beyond an electroporator. The protocol was tailored to ensure that the sequence of the Chlamydomonas genomic DNA flanking the random insertion is consistently obtained in a high proportion of transformants. A detailed protocol is presented to aid in the experimental design and implementation of mutant screens in Chlamydomonas.

Electronic supplementary material
The online version of this article (doi:10.1186/s13007-017-0170-x) contains supplementary material, which is available to authorized users.

Full text

Pollock et al. Plant Methods (2017) 13:22
DOI 10.1186/s13007-017-0170-x
METHODOLOGY
A robust protocol forecient
generation, andgenomic characterization
ofinsertional mutants ofChlamydomonas
reinhardtii
Steve V. Pollock1, Bratati Mukherjee1, Joanna Bajsa‑Hirschel1, Marylou C. Machingura1, Ananya Mukherjee1,
Arthur R. Grossman2 and James V. Moroney1*
Abstract
Background: Random insertional mutagenesis of Chlamydomonas reinhardtii using drug resistance cassettes has
contributed to the generation of tens of thousands of transformants in dozens of labs around the world. In many
instances these insertional mutants have helped elucidate the genetic basis of various physiological processes in this
model organism. Unfortunately, the insertion sites of many interesting mutants are never defined due to experimen‑
tal difficulties in establishing the location of the inserted cassette in the Chlamydomonas genome. It is fairly com‑
mon that several months, or even years of work are conducted with no result. Here we describe a robust method to
identify the location of the inserted DNA cassette in the Chlamydomonas genome.
Results: Insertional mutants were generated using a DNA cassette that confers paromomycin resistance. This
protocol identified the cassette insertion site for greater than 80% of the transformants. In the majority of cases the
insertion event was found to be simple, without large deletions of flanking genomic DNA. Multiple insertions were
observed in less than 10% of recovered transformants.
Conclusion: The method is quick, relatively inexpensive and does not require any special equipment beyond an
electroporator. The protocol was tailored to ensure that the sequence of the Chlamydomonas genomic DNA flanking
the random insertion is consistently obtained in a high proportion of transformants. A detailed protocol is presented
to aid in the experimental design and implementation of mutant screens in Chlamydomonas.
© The Author(s) 2017. This article is distributed under the terms of the Creative Commons Attribution 4.0 International License
(http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium,
provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license,
and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/
publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
Background
Over the past decade, Chlamydomonas reinhardtii (here-
after referred to as Chlamydomonas) has been success-
fully used as a model system to help answer biological
questions related to a wide variety of cellular processes.
With a sequenced genome and a growing experimen-
tal toolbox to facilitate large-scale forward and reverse
genetic studies, this unicellular microalga now provides
an even stronger functional genomics template for the
further dissection of biological processes, and both
metabolic and regulatory pathways. Experimental work
using Chlamydomonas will not only contribute to our
increased understanding of its own physiology and bio-
chemistry, but will continue to reveal the genetic basis
of similar processes in other organisms such as bacteria,
fungi, vascular plants, animals and even humans.
A valuable resource for studying biological processes
is the availability of stable mutations that disrupt key
genes that encode components of those processes. In
Chlamydomonas, insertional mutagenesis has been rou-
tinely used for this purpose, and with great success. A
large number of transformants can be generated using
this technique with the goal of tagging a single func-
tional gene within the nucleus of each transformant.
When transforming nuclear DNA, a short DNA cassette
Open Access
Plant Methods
*Correspondence: btmoro@lsu.edu
1 Department of Biological Sciences, Louisiana State University, Baton
Rouge, LA 70803, USA
Full list of author information is available at the end of the article
Content courtesy of Springer Nature, terms of use apply. Rights reserved.

Page 2 of 9
Pollock et al. Plant Methods (2017) 13:22
consisting of an antibiotic resistance gene marker flanked
by endogenous promoter and terminator sequences is
used to ensure optimum marker gene expression and pre-
vent transcriptional read through. In Chlamydomonas,
the absence of homologous recombination in the nuclear
genome means that insertion of the antibiotic cassette
occurs at random genomic sites. Identification of the
location of the insert in the genome is therefore pivotal
to the success of this method. In forward genetic studies,
the position of the inserted DNA in mutants with pheno-
types of interest would help identify genes disrupted in
the transformants. However, this approach could also be
valuable for reverse genetic screens since there are many
Chlamydomonas genes, both with known and unknown
functions, in which knockouts might not result in readily
discernible phenotypes.
With the current availability of a sequenced and largely
annotated nuclear genome [1, 2], a relatively short
sequence flanking the genomic insert is often provides
enough information to use the genomic database to iden-
tify the disrupted genomic locus. Several techniques were
successfully used in the past to identify insertion sites
within the genome. ese involved modified protocols
for plasmid rescue [3], ermal asymmetric interlaced
PCR or TAIL PCR [4], Restriction enzyme site-directed
amplification PCR or RESDA-PCR [5], 3′-Rapid Amplifi-
cation of cDNA ends or 3′RACE [6] and Site Finding PCR
[7]. Recently, a high throughput Mme1-based Insertion
site Sequencing strategy for Chlamydomonas insertional
mutants, called ChlaMmeSeq [8, 9], was used for the
simultaneous screening of large numbers of mutagenic
insertion sites. Keeping in mind the available technology
used with varying degrees of success in Chlamydomonas,
this study proposes the use of a different protocol aimed
at the successful recovery of genomic regions flank-
ing an insert within transformants generated in large
scale insertional mutagenesis efforts. is protocol is
based on an adaptor-linked PCR method that has been
modified and refined for success with Chlamydomonas.
It is robust, time efficient, and identifies the location of
inserts in the majority of transformants. is method
also detects the number of insertions and their direction,
and any deletions/rearrangements at the insertion site.
Adaptor linked PCR has been used with prokaryotes
[10] and eukaryotes for genome walking, as well as to
identify the location of T-DNA inserts in the Arabidop-
sis genome [11]. is procedure usually involves restric-
tion and blunting of genomic DNA, followed by ligation
of an asymmetric adaptor DNA oligonucleotide that
has been modified to prevent self-ligation. e adap-
tor linked to the ends of genomic DNA provides a tem-
plate for designing primers based on known sequences
of the adaptor and of the inserted cassette for a series of
nested PCRs with varying stringency. ese PCRs pro-
duce DNA fragments that are sequenced and aligned to
the nuclear genome sequence. In this study, the adap-
tor linked PCR method was modified for efficient use in
determining insert locations in Chlamydomonas trans-
formants derived from a large-scale mutagenesis effort.
Several combinations of restriction enzymes and PCR
conditions were tested to provide a robust and relatively
fail-safe method of determining insert location on the
Chlamydomonas nuclear genome.
A detailed protocol is provided to aid researchers
interested in high throughput determination of insert
locations with a population of transformants. Several
recommendations in areas ranging from primer design
to the use of combinations of restriction enzymes that
might further increase the probability of insert recovery
are presented. Many shortcuts have also been suggested
such as one step digestions and ligations, rapid methods
for agarose gel electrophoresis, and the use of shorter
PCR cycles to reduce both time and cost. e execution
of the technique proposed in this study will contribute to
both the generation of genome wide mutant libraries and
the characterization of the insertion sites.
Methods
Generation ofthe insertional DNA fragment
Two methods were used to generate the cassette used
for transformation. One used digestion from the pSL18
plasmid and the other used PCR amplification. e PCR
generated DNA fragment was amplified from vector
pSL72 [12] and transformed into C. reinhardtii strain
D66 (nit2; cw15; mt+) to confer resistance to paromomy-
cin. Primers, RIM-f2 and RIM-r1 (see Table1 for primer
sequences), were used to amplify a DNA fragment with
122bp of the bacterial pBluescript vector, 803bp of the
Chlamydomonas PSAD promoter, 811 bp of the Aph-
VIII coding sequence from Streptomyces rimosus [13]
Table 1 The primers used forthis work
Primer Description Sequence 5′–3′
RIM‑f2 Used to amplify cassette TGT GTG GAA TTG TGA GCG G
RIM‑r1 Used to amplify cassette CTT TCC ATC GGC CCA GCA
RIM 3‑1 3′ insert primer CGG TAT CGG AGG AAA AGC TG
RIM 3‑2 3′ insert primer GCT GTT GGA CGA GTT CTT CTG
RIM 5‑1 5′ insert primer TTC CAA GCG ATC ACC AGC AC
RIM5‑2 5′ insert primer GCT GGC ACG AGT ACG GGT TG
RIM5‑4 5′ insert primer AGC TTT TGT TCC C TT TAG TG
AP1 Adaptor primer GTA ATA CGA CTC ACT ATA GAG T
AP2 Adaptor primer ACT ATA GAG TAC GCG TGG T
RX1 5′ insert primer pSL18 GCC CTC ATA GCC CGC CAA ATC AG
RX2 5′ insert primer pSL18 AAG CCG ATA AAC ACC AGC CC
Content courtesy of Springer Nature, terms of use apply. Rights reserved.

Page 3 of 9
Pollock et al. Plant Methods (2017) 13:22
and 58bp of the 5′ end of the second intron of the CYC6
gene of Chlamydmonas (Fig.1a). e Expand Long Tem-
plate PCR System (Roche; CAT. #11681834001), a mix-
ture of thermostable TAQ DNA polymerase and Tgo (a
proofreading DNA polymerase) and approximately 10ng
of the pSL72 plasmid template were used to amplify the
1812bp DNA fragment. e composition of a single PCR
reaction is shown in Table 2. e temperature cycling
protocol is shown in Fig.2. Alternatively, the cassette was
obtained by restriction digestion of the pSL18 plasmid.
XhoI and NheI (NEB) were used to digest the plasmid.
e resulting fragment contained the HSP70-RbcS2 dual
promoter followed by the paromomycin gene and the
RBCS2 terminator (Fig.1b).
Preparation ofthe DNA insert
After electrophoresis in an 0.8% (W/V) agarose gel, the
DNA fragments were purified from the gel by excising
the 1812bp fragment (PCR generated) or the 1813bp
fragment (restriction generated) with a razor blade (vis-
ualized with ethidium bromide and a low energy UV
light). DNA from the gel slices were purified using a gel
extraction kit (Qiagen). e purified DNA was filter steri-
lized and quantified by comparing the fluorescence of
the DNA to that of a known DNA mass ladder (HindIII
digested lamda DNA (New England Biolabs); or a 1 Kb
ladder (New England Biolabs).
Fig. 1 The paromomycin cassette. Two different methods were used to generate the paromomycin cassette used in these experiments. a PCR gen‑
erated cassette. The paromomycin resistance cassette used to generate paromomycin resistant strains, and the primers used to amplify the cassette
(RIM‑f2, and RIM‑r1) from pSL72. The other primers indicated in the figure were used to amplify genomic DNA flanking the site of the insertion. b
Restriction digested generated cassette. The paromomycin cassette from the pSL18 plasmid
Table 2 Composition of the 50 µL PCR reaction mixture
used toamplify the 1812bp insertional DNA fragment
a Containing approx. 10ng plasmid DNA
Component Volume
dH2O 37.5 µL
10× polymerase buffer #3 5 µL
2.5 mM dNTP mix 4 µL
RIM‑f2 (20 µM) 1 µL
RIM‑r1 (20 µM) 1 µL
pSL72 plasmid template 1 µLa
Polymerase mix (5 U µL−1) 0.5 µL
Fig. 2 The temperature cycling protocol used to amplify the inser‑
tional DNA fragment
Content courtesy of Springer Nature, terms of use apply. Rights reserved.

Page 4 of 9
Pollock et al. Plant Methods (2017) 13:22
Preparing chlamydomonas cells fortransformation
500mL of TAP medium [14] was inoculated with a 4mm
spherical scraping (a small green pea volume equivalent)
of D66 cells from a TAP agar plate. When cells reached
a density of approximately 3×106cellsmL−1 they were
harvested by centrifugation at 2500×g and resuspended
in a final volume of 3.5–4.0mL TAP containing 60mM
sorbitol to achieve a final cell concentration of 2–4×108
cells per mL. Resuspended cells were transferred to a
15mL Corning centrifuge tube (Corning #430790) and
placed on ice. Electroporation cuvettes (Bio-Rad #165-
2091) with a gap-width of 0.4 cm were used for elec-
troporation. 250µL of cells was transferred to a sterile
cuvette with no added DNA, labeled “No DNA control”,
and placed on ice. Approximately 2.5µg of the prepared
insert DNA fragment was added to the remaining 3.25–
3.75 mL of suspended cells, gently inverted 5–6 times
to make the DNA/cell mixture homogeneous and then
placed on ice. 250µL aliquots of the DNA/cell suspen-
sion were added to the sterile electroporation cuvettes,
yielding approximately 14 transformation reactions. All
cuvettes were placed on ice for 10–20min before elec-
troporation. is step is critical in obtaining a pulse rate
that does not lyse the cells (see below). Care must be
taken to ensure the cells do not warm up before the pulse
is generated.
Electroporation ofchlamydomonas
As stated above, 250 µL containing between 0.5 ×108
and 1×108cells with 180ng of the paromomycin cas-
sette were electroporated using the Bio-Rad Gene Pul-
ser II system, as modified from the method reported by
Shimogawara et al. [15]. One significant modification
was that no carrier DNA was added. A voltage setting of
0.8kV, a capacitor setting of 25µF, and no shunt resistor,
was used for electroporation. e measured pulse time,
an excellent predictor of the success of the electropora-
tion transformation, generally ranged from 10 to 13ms. If
the pulse time was shorter, the transformation efficiency
decreased approximately 100-fold as cells were lysed
during the pulse. e cuvettes were placed at room tem-
perature for 5min following electroporation and prior to
transfer to overnight recovery medium.
Overnight recovery ofelectroporated cells
Within 30min of electroporation, the mixture from each
cuvette was transferred using a 200µL wide orifice pipet
tip (E&K #3502-R96S) (to prevent shearing of the cells)
to 10 mL of TAP medium containing 60mM sorbitol
in a 15mL Corning centrifuge tube (Corning #430790).
e tubes were placed in low light (10–20 µmol pho-
tons m−2s−1), gently rocked to keep the cells suspended,
and allowed to recover overnight (12–16 h). Used
electroporation cuvettes were rinsed with water and
stored in 100% ethanol until required for another round
of transformations. As long as the cuvettes did not crack
they were reused several times. Cuvettes were dried in a
sterile hood immediately before use.
Plating recovered electroporated cells onselective
medium
Petri dishes with solid TAP medium containing 1.5% agar
(W/V) and 7.5µgmL−1 of paromomycin sulfate (Sigma
# P5057; stock of 100mgmL−1 dissolved in dH2O, filter
sterilized, and frozen) were prepared one day in advance
of plating. e 10mL of recovered cells were harvested
by centrifugation in an IEC swing-out clinical centri-
fuge at maximum speed for 1 min and resuspended in
80–100µL of TAP medium. e entire mixture was gen-
tly spread, using a bent glass rod, on a single TAP plus
paromomycin plate and allowed to dry in a transfer hood.
Dried plates were placed in moderate light (50–80µmol
photons m−2s−1). Within 2days of plating the majority of
the cells began to die and after 4days small colonies were
noted under a dissecting microscope. After 1 week the
colonies were large enough to pick with sterile pointed
toothpicks onto a screening plate using a 10× 10 grid
(Fig.3). e typical yield of paromomycin resistant trans-
formants varied from 150 to 300 colonies per plate. is
corresponds to roughly 4 transformants per 106 cells.
Once accustomed to picking colonies, a single researcher
can “easily” pick 500 colonies in 2h. A self-closing pair of
tweezers made holding toothpicks more comfortable and
Fig. 3 The 10 × 10 grid template used to plate the paromomycin
transformants. The D66 squares are the untransformed parental
control blocks
Content courtesy of Springer Nature, terms of use apply. Rights reserved.

Page 5 of 9
Pollock et al. Plant Methods (2017) 13:22
allowed the user to use both ends of the toothpicks. Used
toothpicks were reused after autoclaving and drying.
Genomic DNA preparation
Total DNA was isolated from 50 mL of mutant cells
grown in TAP medium under continuous low light (50–
80µmol photons m−2s−1) according to Newman etal.
[16] with several modifications. Briefly, cells were pelleted
by centrifugation at 2500×g in 50mL sterile centrifuge
tubes (Corning #430828) and resuspended in 400µL of
dH20 in two 1.5mL Eppendorf tubes and 800µL disrup-
tion buffer containing SDS was added (2% SDS, 400mM
NaCl, 40mM EDTA, 100mM Tris–HCl, pH 8.0). e
nucleic acids were extracted three times using a phenol/
chloroform/isoamyl alcohol mixture until the interface
of the inorganic and organic layers contained no residual
protein residue. e aqueous phase was then extracted a
final time with chloroform. Nucleic acids were then pre-
cipitated with two volumes of ethanol and washed twice
with 70% ethanol. e pellet was air-dried for 10min and
dissolved in 100µL of TE (10 mM TRIS, pH7.5, 1mM
EDTA). See Additional file1 for a more detailed descrip-
tion of the DNA preparation protocol.
Preparation ofblunt‑ended restriction digest fragments
ofgenomic DNA
200ng of genomic DNA was digested with a mixture of
AleI (10 units), NaeI (10 units), PmlI (10 units) and PvuII
(1 unit) restriction endonucleases (New England Biolabs)
in a volume of 100 µL in NEB buffer 2 supplemented
with 100µgmL−1 BSA. e reactions were incubated at
37°C for 16–18h. ese four restriction endonucleases
were chosen because they do not recognize sequences
in the insertion DNA fragment and because they recog-
nize sequences that occur, on average, every 200bps in
the C. reinhardtii genome, creating blunt-ended frag-
ments. e last hour of the reaction was supplemented
with 1µL RNAse to degrade RNA. After the incubation,
5 µL of digested genomic DNA was separated by aga-
rose gel electrophoresis to verify that the digestions were
complete, which was observable as a smear of DNA from
approximately 1 to 6kb on a 1% agarose gel. Digested
DNA was extracted once with an equal volume (95µL)
of phenol:chloroform:isoamyl alcohol (25:24:1 v/v), and
once with chloroform:isoamylalcohol (24:1v/v), precipi-
tated with three volumes (285mL) of ethanol, the pellet
washed once with ice-cold 80% (V/V) ethanol, air-dried
for 10min and then resuspended in 20µL of TE.
Adaptor preparation
A blunt-ended adaptor consisting of a 48bp DNA oligo,
designated plus strand, and a 10bp oligonucleotide, des-
ignated negative strand, were procured from Integrated
DNA Technologies. e plus strand, 5′-GTA ATA CGA
CTC ACT ATA GAG TAC GCG TGG TCG ACG GCC
CGG GCT GGT-3′, was procured (250nmol level) and
HPLC purified by the manufacturer. e minus strand,
5′-ACC AGC CCG G-3′, was procured at the (100nmol
level), with a 3′ C3 spacer to prevent polymerase exten-
sion, and 5′ phosphorylation to permit ligation, and
HPLC purified. e two strands were each dissolved in
STE (10mM Tris (pH 8.0), 50mM NaCl, 1mM EDTA)
at a concentration of 50µM and 25µL of each strand was
mixed together and placed in a thermocycler at 95 °C.
e thermocycler was programmed to gradually cool to
4°C over a period of approximately 3h to allow the two
strands to anneal to form a double stranded asymmetric
blunt-ended adaptor (Fig.4). e resulting adaptor is sta-
ble and can be stored at 4°C or frozen. To prevent dena-
turation of the adaptor, it was kept cool during handling.
Ligation ofthe adaptor tothe digested DNA
e adaptor was ligated to the digested genomic DNA
overnight (16–20 h) at 16 °C. e ligation reaction
Fig. 4 The adaptor used to obtain the flanking DNA. The blunt‑ended asymmetric adaptor, consisting of a 48 bp positive strand (+) and a 10 bp
negative strand (−), and the two adaptor primers (AP1 and AP2) aligned to depict where they will bind after the negative strand of the adaptor is
extended in the first round of the primary PCR reaction
Content courtesy of Springer Nature, terms of use apply. Rights reserved.

Page 6 of 9
Pollock et al. Plant Methods (2017) 13:22
consisted of 2.0µL of adaptor, 4µL of digested genomic
DNA, 2 µL of 10× New England Biolabs ligase buffer,
1µL of T4 DNA ligase (NEB #M0202S). e reaction was
stopped by incubating the mixture at 80°C for 20min.
Seventy µL of TE was added to the reaction before use as
template during the PCR amplification.
Alternative one‑step restriction digest andligation
reaction
Alternatively, the genomic DNA was restricted and
ligated to the adaptor in a one-step reaction. e one step
reaction was performed in the restriction enzyme buffer
(NEB#2) with the addition of 10µM ATP. e reaction
was performed overnight at room temperature.
PCR amplication fromthe insertion tothe anking
adaptor
Primary and nested PCR reactions were used to amplify
the genomic DNA flanking the insertion of the paromo-
mycin resistance cassette. By using primer sets directed
out from the 5′ or 3′ ends of the insert, it was possible
to amplify DNA flanking both sides of the insert (Fig.5).
e primary reaction utilized an insert specific primer
(RIM3-1; or RIM5-1) and an adaptor primer (AP1) (see
Table1 for lists of primers). RIM3-1 and RIM5-1 were
used to amplify the 3′ and 5′ flanking DNA respectively
(Fig. 1). To enrich for amplification from the paromo-
mycin insertion the AP1 primer binding site was only
generated after the RIM3-1 primer extended the 10bp
strand of the adaptor to yield an adaptor sequence
for AP1 primer binding. A touch-down PCR protocol
using Expand Long Template PCR System (Roche; CAT.
#11681834001) was utilized for the primary and nested
PCR reactions (Fig.6). e primary reaction was diluted
50-fold by resuspending 1 µL of the primary reaction
in 49µL of dH2O and vortexed to mix. e nested PCR
reaction was then performed using the nested primers
RIM3-2 (or RIM 5-2) and the adaptor primer AP2 using
the same cycling parameters used in the primary reac-
tion. In some instances the use of an additional nested
PCR reaction using RIM 5-4 was necessary to obtain
a 5′ flanking DNA fragment. e RIM5-4 primer was
designed to bind to the portion of the cassette arising
from pBluescript. All DNA fragments amplified by the
above procedure were sequenced using the last insertion
specific primer (RIM 3-2, or RIM5-4).
Results anddiscussion
Adaptor PCR
In this investigation, 30,000 insertional mutants were
selected following transformation with the paromomy-
cin resistance conferring cassette. After this selection,
colonies were screened for growth on high and low CO2
and 211 colonies showing a growth deficiency only under
low CO2 conditions were chosen for further analysis.
Adaptor PCR was performed on the genomic DNA of
the mutants. e majority of the mutants yielded a sin-
gle PCR product, but some also produced two or more
products. In most cases, the length of the flanking DNA
ranged from 150 bp to 1200bp while some fragments
>2000 bps were obtained. A representative agarose gel
is shown in Fig.7. Fragments were excised from the gels
and sequenced. e majority of the sequenced fragments
contained the 3′ end of the paromomycin resistance cas-
sette followed by Chlamydomonas genomic DNA. Some
PCR products were too short to map accurately to the
Chlamydomonas genome (for instance, fragments less
than 15 nucleotides in length), and some contained only
the paromomycin vector sequence. However, we were
able to map over 74% of the sequences (156 out of 211
independent inserts) to the Chlamydomonas genome.
From the 156 colonies where the DNA flanking the
insert was identified, 36 colonies were chosen for further
molecular analysis. is analysis included mapping both
ends of the insert as well as checking for DNA deletions
in the regions flanking the paromomycin cassette. When
both ends of the insert were mapped we found that 75%
of the mapped inserts (27/36) represented simple inser-
tions of the resistance cassette resulting in no genomic
deletion of>10bp. Of the remaining 25% of the colonies
(9/36), we were unable to map the 3′ end of the insert to
the same genomic region identified at the 5′ end. us, in
some instances a Chlamydomomas genomic fragment is
inserted between the cassette and the true genomic loca-
tion. is result is similar to that of Zhang etal. [8] who
mapped inserts using the ChlaMmeSeq method. Overall,
the results show that 75% of the insertional mutants gen-
erated by our mutagenesis resulted in a simple insertion
that sometimes had a small DNA deletion.
is report details a method that enables researchers to
generate insertional mutants and to successfully identify
the location of the paromomycin resistance cassette in
the Chlamydomonas genome over 75% of the time. e
method is rapid, fairly inexpensive and does not require
exotic equipment, except for the electroporator. is
method differs significantly from other methods which
primarily use TAIL PCR or a modification of that method
[4, 5]. First, the transformation method described in
this report was electroporation versus glass beads, to
reduce genomic deletions. Secondly, a DNA cassette was
used containing only the paromomycin resistance gene
instead of a plasmid [4, 5]. e use of a cassette makes
it easier to identify the ends of the inserted DNA and
makes the adaptor method a viable choice for research-
ers. In addition, a combination of restriction enzymes
is described that enhances the likelihood of cutting the
Content courtesy of Springer Nature, terms of use apply. Rights reserved.

Page 7 of 9
Pollock et al. Plant Methods (2017) 13:22
flanking genomic DNA within 1000 bp of the insert,
increasing the success of obtaining a PCR fragment long
enough to identify the site of insertion. Finally, adap-
tor PCR was used instead of TAIL PCR, which offers an
additional tool to use if a researcher is having trouble
with one method. Recently, a library of Chlamydomonas
insertional mutants became available to researchers [9].
is major advance will allow researchers to order Chla-
mydomonas strains containing an insert in specific genes.
However, for gene discovery, scientists will still need to
generate and screen new insertional mutants in their
own laboratories. After conducting these screens and
selecting strains with the desired phenotype, research-
ers need to be able to identify the disrupted gene. Fur-
thermore, the new insertion library is not complete (less
than 40% of the genes covered by more than one allele,
and only a small number of the mutants have been vali-
dated). Finally, the method for generating insertional
mutants described here will result in a high percentage
of the colonies having single, simple insertions with few
large genomic deletion. is transformation procedure
also yields a relatively low number of strains with multi-
ple DNA insertions. We found that multiple inserts were
present in 11 out of the 156 mutants (~6%), although
some insertions will not be detected using this method.
Zhang etal. [8] observed that about 15% of the paromo-
mycin resistant transformants had more than one insert
when using a similar transformation protocol. We also
observed very few deletions of genomic DNA flanking
the inserts. One disadvantage of the glass bead trans-
formation method is that it sometimes results in large
DNA deletions resulting in the loss of more than one
gene, making the results much harder to analyze [17, 18].
e electroporation method appears to be less likely to
cause these large deletions although deletions have been
reported using this method also [19, 20]. While we did
not find large deletions in the transformants that we
characterized, we were unable to recover the other side of
the insert in 25% of the colonies, which could indicate the
occurrence of large deletions or rearrangements.
Fig. 5 The adaptor PCR method. A flow‑chart depicting the adaptor‑mediated PCR method to obtain DNA sequence flanking the insertion of the
paromomycin resistance cassette (AphVIII). The bold text in the final sequence highlights the 3′ end of the insertion DNA, and the bold/italicized text
depicts the 48 bp sequence that is sometimes present in shorter DNA fragments
Content courtesy of Springer Nature, terms of use apply. Rights reserved.

Page 8 of 9
Pollock et al. Plant Methods (2017) 13:22
Cosegregation ofthe DNA insert withthe phenotype being
studied
For any study using insertional mutagenesis, cosegrega-
tion of the insert with the desired phenotype must be
genetically demonstrated. Genetic linkage was inves-
tigated in 15 strains generated using this method and
paromomycin resistance cosegregated with the SLC
phenotype about 40% of the time (6 of 15 transfor-
mants). Cells with an SLC phenotype grow normally
on elevated CO2 concentrations but more slowly than
wild-type cells at low CO2 concentrations. is rate of
cosegregation is similar rates reported by others [4].
Since random insertion involves double strand DNA
breaks the generation of insertions and point mutations
is likely. Clearly it remains essential that a genetic analy-
sis of any interesting insertional mutant be done before
proceeding with a complete physiological characteri-
zation of the mutant. Tetrad analysis was used in this
study but random spore analysis could also be used. It
is also critical to demonstrate rescue of the mutant phe-
notype by introduction of a wild type copy of the dis-
rupted locus.
Earlier successes
Earlier versions of the transformation procedure
described here were used to generate and characterize
insertional mutants in two large scale experiments. In
one study, over 30,000 insertional mutants were screened
for aberrant responses to sulfur limitation (SAC) [21].
In a separate investigation, again over 30,000 insertional
mutants were generated and screened for a ‘sick in low
carbon dioxide’ phenotype (SLC). In some cases, mutants
showing the desired phenotype were subjected to adaptor
PCR to determine the genomic location of the paromo-
mycin resistance cassette in their genomes. In the study
in which the cells were screened for aberrant responses
to sulfur limitation, a number of novels genes were dis-
covered, including proteins involved in the responses of
Chlamydomonas to sulfur deprivation [19, 21]. In the
screen for mutants unable to grow photoautotrophically
on low CO2, insertions in CIA6 [22], bestrophin, MITC11
and LCI9 [23] andCIA8 (Machingura, Bajsa-Hirschel and
Moroney, unpublished) were identified using the forward
genetics approach described by González-Ballester etal.
[5]. No large deletions were observed in these studies.
Fig. 6 The touch‑down PCR protocol. The touch‑down temperature
cycling protocol used to amplify DNA flanking the insertional DNA
fragment
Fig. 7 Representative results using this adaptor PCR method. Repre‑
sentative results from several insertional mutants (A1–A24) following
the described protocol. Agarose gel electrophoresis was used to
visualize the products of the primary and secondary PCR reactions
respectively for each mutant. A diagnostic step‑down in fragment
size was indicative of a positive result as the nested primers amplified
the target DNA from the mutants
Content courtesy of Springer Nature, terms of use apply. Rights reserved.

Page 9 of 9
Pollock et al. Plant Methods (2017) 13:22
Conclusion
In this communication we have detailed a method to
generate insertional mutants that will have mostly single
simple DNA inserts. We have described an adaptor-PCR
based method that reliably can identify the location of the
cassette in Chlamydomonas. Using this method, we were
able to identify the genomic DNA flanking the insertion
over 75% of the time. e combination of employing the
electroporation method to generate insertional mutants
in conjunction with the adaptor method should provide
researchers using Chlamydomonas an excellent chance
to quickly generate and characterize useful insertional
mutant strains at a relatively low cost.
Authors’ contributions
SVP, ARG and JVM designed the method, all authors conducted the experi‑
ments and helped write the manuscript. All authors read and approved the
final manuscript.
Author details
1 Department of Biological Sciences, Louisiana State University, Baton Rouge,
LA 70803, USA. 2 Department of Plant Biology, Carnegie Institution for Science,
Stanford, CA 94305, USA.
Acknowledgements
The authors thank Susan Laborde for excellent technical assistance.
Competing interests
The authors declare that they have no competing interests.
Consent for publication
All authors have consented for publication.
Funding
Supported by NSF Award IOS1146597 and subcontract from the University of
Illinois to JVM and NSF Award MCB 0951094 to ARG.
Received: 19 October 2016 Accepted: 22 March 2017
References
1. Merchant SS, Prochnik SE, Vallon O, Harris EH, et al. The Chlamydomonas
genome reveals the evolution of key animal and plant functions. Science.
2007;318:245–50.
2. Blaby IK, Blaby‑Haas CE, Tourasse N, Hom EFY, Lopez D, Aksoy M, Gross‑
man A, Umen J, Dutcher S, Porter M, King S, Witman GB, Stanke M, Harris
EH, Goodstein D, Grimwood J, Schmutz J, Vallon O, Merchant SS, Prochnik
S. The Chlamydomonas genome project: a decade on. Trends Plant Sci.
2014;19:672–80.
3. Tam LW, Lefebvre PA. Cloning of flagellar genes in Chlamydomonas
reinhardtii by DNA insertional mutagenesis. Genetics. 1993;135:375–84.
4. Dent RM, Haglund CM, Chin BL, Kobayashi MC, Niyogi KK. Functional
genomics of eukaryotic photosynthesis using insertional mutagenesis of
Chlamydomonas reinhardtii. Plant Physiol. 2005;137:545–56.
Additional le
Additional le1. A simple Protocol for obtaining Chlamydomonas
genomic DNA.
5. González‑Ballester D, Montaigu A, Galván A, Fernández E. Restriction
enzyme site‑directed amplification PCR: a tool to identify regions flanking
a marker DNA. Anal Biochem. 2005;340:330–5.
6. Meslet‑Cladière L, Vallon O. A new method to identify flanking sequence
tags in Chlamydomonas using 3′‑RACE. Plant Methods. 2012;8:1.
7. Li X, Moelleringa ER, Liu B, Johnny C, Fedewac M, Sears BB, Kuoc MH, Ben‑
ning C. A galactoglycerolipid lipase is required for triacylglycerol accu‑
mulation and survival following nitrogen deprivation in Chlamydomonas
reinhardtii. Plant Cell. 2012;24:4670–86.
8. Zhang RR, Patena W, Armbruster U, Gang SS, Blum SR, Jonikas MC.
High‑throughput genotyping of green algal mutants reveals random
distribution of mutagenic insertion sites and endonucleolytic cleavage of
transforming DNA. Plant Cell. 2014;26:1398–409.
9. Li X, Zhang R, Patena W, Gang SS, Blum SR, Ivanova N, Yue R, Robertson
JM, Lefebvre PA, Fitz‑Gibbon ST, Grossman AR, Jonikas MC. An indexed,
mapped mutant library enables reverse genetics studies of biological
processes in Chlamydomonas reinhardtii. Plant Cell. 2016;28:367–87.
10. Wang S, He J, Cui Z, Li S. Self‑formed adaptor PCR: a simple and
efficient method for chromosome walking. Appl Environ Microbiol.
2007;73:5048–51.
11. O’Malley RC, Alonso JM, Kim CJ, Leisse TJ, Ecker JR. An adapter ligation‑
mediated PCR method for high‑throughput mapping of T‑DNA inserts in
the Arabidopsis genome. Nat Protoc. 2007;2:2910–7.
12. Pollock SV, Colombo SL, Prout DL Jr, Godfrey AC, Moroney JV. Rubisco
activase is required for optimal photosynthesis in the green alga
Chlamydomonas reinhardtii in a low CO2 atmosphere. Plant Physiol.
2003;133:1854–61.
13. Sizova I, Fuhrmann M, Hegemann P. A Streptomyces rimosus aphVIII gene
coding for a new type phosphotransferase provides stable antibiotic
resistance to Chlamydomonas reinhardtii. Gene. 2001;277:221–9.
14. Sueoka N. Mitotic replication of deoxyribonucleic acid in Chlamydomonas
reinhardtii. Proc Natl Acad Sci USA. 1960;46:83–91.
15. Shimogawara K, Fujiwara S, Grossman AR, Usuda H. High efficiency trans‑
formation of Chlamydomonas reinhardtii by electroporation. Genetics.
1998;148:1821–8.
16. Newman SM, Boynton JE, Gillham NW, Randolph‑Anderson BL, Johnson
AM, Harris EH. Transformation of chloroplast ribosomal RNA in Chla‑
mydomonas: molecular and genetic characterization of integration
events. Genetics. 1990;126:875–88.
17. Dent RM, Sharifi MN, Malnoë A, Haglund C, Calderon RH, Wakao S, Niyogi
KK. Large‑scale insertional mutagenesis of Chlamydomonas supports
phylogenomic functional prediction of photosynthetic genes and analy‑
sis of classical acetate‑requiring mutants. Plant J. 2015;82:337–51.
18. Wang Y, Spalding MH. An inorganic carbon transport system responsible
for acclimation specific to air levels of CO2 in Chlamydomonas reinhardtii.
Proc Natl Acad Sci USA. 2006;103:10110–5.
19. Aksoy M, Pootakham W, Pollock SV, Moseley JL, González‑Ballester
D, Grossman AR. Tiered regulation of sulfur deprivation responses in
Chlamydomonas reinhardtii and identification of an associated regulatory
factor. Plant Physiol. 2013;162:195–211.
20. Aksoy M, Pootakham W, Grossman AR. Critical function of a Chla-
mydomonas reinhardtii putative polyphosphate polymerase subunit
during nutrient deprivation. Plant Cell. 2014;26:4214–29.
21. Gonzalez‑Ballester D, Pollock SV, Pootakham W, Grossman AR. The central
role of a SNRK2 kinase in sulfur deprivation responses. Plant Physiol.
2008;147:216–27.
22. Ma Y, Pollock SV, Xiao Y, Cunnusamy K, Moroney JV. Identification of a
novel gene required for normal pyrenoid formation in Chlamydomonas
reinhardtii. Plant Physiol. 2011;156:884–96.
23. Jungnick N, Ma Y, Mukherjee B, Cronan JC, Speed DJ, Laborde SM,
Longstreth DJ, Moroney JV. The carbon concentrating mechanism in
Chlamydomonas reinhardtii—finding the missing pieces. Photosynth Res.
2014;121:159–73.
Content courtesy of Springer Nature, terms of use apply. Rights reserved.

1.
2.
3.
4.
5.
6.
Terms and Conditions
Springer Nature journal content, brought to you courtesy of Springer Nature Customer Service Center GmbH (“Springer Nature”).
Springer Nature supports a reasonable amount of sharing of research papers by authors, subscribers and authorised users (“Users”), for small-
scale personal, non-commercial use provided that all copyright, trade and service marks and other proprietary notices are maintained. By
accessing, sharing, receiving or otherwise using the Springer Nature journal content you agree to these terms of use (“Terms”). For these
purposes, Springer Nature considers academic use (by researchers and students) to be non-commercial.
These Terms are supplementary and will apply in addition to any applicable website terms and conditions, a relevant site licence or a personal
subscription. These Terms will prevail over any conflict or ambiguity with regards to the relevant terms, a site licence or a personal subscription
(to the extent of the conflict or ambiguity only). For Creative Commons-licensed articles, the terms of the Creative Commons license used will
apply.
We collect and use personal data to provide access to the Springer Nature journal content. We may also use these personal data internally within
ResearchGate and Springer Nature and as agreed share it, in an anonymised way, for purposes of tracking, analysis and reporting. We will not
otherwise disclose your personal data outside the ResearchGate or the Springer Nature group of companies unless we have your permission as
detailed in the Privacy Policy.
While Users may use the Springer Nature journal content for small scale, personal non-commercial use, it is important to note that Users may
not:
use such content for the purpose of providing other users with access on a regular or large scale basis or as a means to circumvent access
control;
use such content where to do so would be considered a criminal or statutory offence in any jurisdiction, or gives rise to civil liability, or is
otherwise unlawful;
falsely or misleadingly imply or suggest endorsement, approval , sponsorship, or association unless explicitly agreed to by Springer Nature in
writing;
use bots or other automated methods to access the content or redirect messages
override any security feature or exclusionary protocol; or
share the content in order to create substitute for Springer Nature products or services or a systematic database of Springer Nature journal
content.
In line with the restriction against commercial use, Springer Nature does not permit the creation of a product or service that creates revenue,
royalties, rent or income from our content or its inclusion as part of a paid for service or for other commercial gain. Springer Nature journal
content cannot be used for inter-library loans and librarians may not upload Springer Nature journal content on a large scale into their, or any
other, institutional repository.
These terms of use are reviewed regularly and may be amended at any time. Springer Nature is not obligated to publish any information or
content on this website and may remove it or features or functionality at our sole discretion, at any time with or without notice. Springer Nature
may revoke this licence to you at any time and remove access to any copies of the Springer Nature journal content which have been saved.
To the fullest extent permitted by law, Springer Nature makes no warranties, representations or guarantees to Users, either express or implied
with respect to the Springer nature journal content and all parties disclaim and waive any implied warranties or warranties imposed by law,
including merchantability or fitness for any particular purpose.
Please note that these rights do not automatically extend to content, data or other material published by Springer Nature that may be licensed
from third parties.
If you would like to use or distribute our Springer Nature journal content to a wider audience or on a regular basis or in any other manner not
expressly permitted by these Terms, please contact Springer Nature at
onlineservice@springernature.com