Extraction of high‐quality, high‐molecular‐weight DNA depends heavily on cell homogenization methods in green microalgae

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Chlorophyte microalgae are important primary producers present

in virtually every photic habitat, including marine, freshwater, hy-

dro-terrestrial, and terrestrial environments (Domozych et al.,

2012). Microalgae have developed physiological and morpho-

logical adaptations to survive in the low humidity, high light, and

uctuating temperatures characteristic of terrestrial environments,

including those of extreme habitats such as deserts, alpine areas,

and polar regions. is naturally occurring diversity, together with

suitability for year-round culturing, has placed green microalgae at

the forefront of applied research (Metting, 1996) in elds includ-

ing bioremediation (Ji etal., 2013), CO

sequestration (Cheah etal.,

2015), heavy metal accumulation (Peña-Castro etal., 2004), biofuels

(Brennan and Owende, 2010), biohydrogen (Nagarajan etal., 2017),

fertilizers (Renuka etal., 2018), and high-value food supplements

and cosmetics (Borowitzka, 2013).

Applied research on microalgae has beneted from our in-

creased knowledge of algal genomics, beginning with the publica-

tion of the rst annotated green algal genomes of Ostreococcus tauri

C. Courties & M.-J. Chrétiennot-Dinet (Derelle etal., 2006) and

Chlamydomonas reinhardtii P. A. Dang. (Merchant etal., 2007) just

over a decade ago. Since then, more than 114 green algal genomes

have become available in the National Center for Biotechnology

Information (NCBI) assembly database (https://www.ncbi.nlm.

nih.gov/assem bly/ query “Chlorophyta” [Organism], consulted

February 2020). is rapid growth was possible because of the

reduction in costs and the development of third-generation

high-throughput sequencing technologies, such as SMRT sequenc-

ing (PacBio, Pacic Biosciences, Menlo Park, California, USA)

and nanopore (Oxford Nanopore Technologies, Oxford, United

Kingdom). ese long-read technologies bypass several of the chal-

lenges in assembling green algal genomes, including co-occurring

bacterial sequences; the presence of nuclear, mitochondrial, and

chloroplast DNA; and high GC content (Blaby etal., 2014). Instead

of relying on assemblies, long reads span through repetitive regions

and low-complexity genomic regions. Long reads, which are able

to resolve complete bacterial genomes in a few reads, are also es-

sential for identifying horizontal gene transfer events in eukaryote

genomes.

published by Wiley Periodicals, Inc. on behalf of the Botanical Society of America. This is an open access article under the terms of the Creative Commons

Attribution-NonCommercial License, which permits use, distribution and reproduction in any medium, provided the original work is properly cited and is

not used for commercial purposes.

Extraction of high-quality, high-molecular-weight DNA

depends heavily on cell homogenization methods in green

microalgae

Jordan R. Stark

, Zoe G. Cardon

, and Elena L. Peredo

1,2

PROTOCOL NOTE

Manuscript received 30 May 2019; revision accepted

8 November 2019.

Ecosystems Center,Marine Biological Laboratory, 7 MBL Street,

Woods Hole, Massachusetts 02543, USA

Author for correspondence: el[email protected]

Citation: Stark, J. R., Z. G. Cardon, and E. L. Peredo. 2020.

Extraction of high-quality, high-molecular-weight DNA depends

heavily on cell homogenization methods in green microalgae.

Applications in Plant Sciences 8(3): e11333.

doi:10.1002/aps3.11333

PREMISE: New sequencing technologies have facilitated genomic studies in green microalgae;

however, extracting high-quality DNA is often a bottleneck for long-read sequencing.

METHODS AND RESULTS: Here, we present a low-cost, highly transferrable method for the

extraction of high-molecular-weight (HMW), high-purity DNA from microalgae. We rst

determined the eect of sample preparation on DNA quality using three homogenization

methods: manual grinding using a mini-pestle, automatic grinding using a vortex adapter,

and grinding in liquid nitrogen. We demonstrated the versatility of grinding in liquid nitrogen

followed by a modied cetyltrimethylammonium bromide (CTAB) extraction across a suite

of aquatic- and desert-evolved algal taxa. Finally, we tested the protocol's robustness by

doubling the input material to increase yield, producing per sample up to 20 μg of high-

purity DNA longer than 21.2 kbp.

CONCLUSIONS: All homogenization methods produced DNA within acceptable parameters

for purity, but only liquid nitrogen grinding resulted in HMW DNA. The optimization of cell

lysis while minimizing DNA shearing is therefore crucial for the isolation of DNA for long-read

genomic sequencing because template DNA length strongly aects read output and length.

KEY WORDS DNA integrity; long-read sequencing; modied CTAB extraction;

Scenedesmaceae.

Applications in Plant Sciences 2020 8(3): e11333 Stark etal.—High-molecular-weight DNA extraction in microalgae • 2 of 7

Long-read sequencing technologies require large quanti-

ties (1 μg to 15 μg, depending on the platform and desired read

length; https://nanop orete ch.com/produ cts/kits, https://www.

pacb.com/wp-conte nt/uploa ds/SMRTb ell-Libra ry-Prepa ratio n-

for-High-Fidel ity-Long-Read-Seque ncing -Custo mer-Train ing.

pdf) of high-purity, high-molecular-weight (HMW) DNA (Rhoads

and Au, 2015). ese concentrations of HMW DNA can be partic-

ularly challenging to obtain from green microalgae. Microalgal cells

are usually small (oen <10 μm), have rigid cell walls, and are rich

in compounds such as chlorophyll a and b, xanthophylls, beta caro-

tene, starch, and cellulose (Lewis and McCourt, 2004), which deeply

inuence the DNA extraction process, aecting cell lysis and down-

stream applications such as PCR amplication (Eland etal., 2012;

Greco etal., 2014). e extraction of DNA from terrestrial algae,

and especially desert-evolved taxa, is notoriously dicult, likely due

to the development of enlarged cell walls during their adaptation to

terrestrial environments (Cardon etal., 2008).

Traditionally, methods to improve the quality of extracted ge-

nomic DNA have focused on purity and yield, as these parameters

have the most impact in the success of downstream applications

(hybridization, PCR, activities of restriction enzymes). e purity

of samples can be increased by ne-tuning extraction protocols

based on the cetyltrimethylammonium bromide (CTAB) extraction

method (Doyle and Doyle, 1987) or by selecting species-appropriate

extraction buers (Tear etal., 2013). Several commercially available

kits using proprietary buers or columns have also been developed

to address the diculty in isolating high-purity DNA from plants

including green microalgae (Eland etal., 2012). Yields can be in-

creased by using maxi-prep approaches, by modifying the amount

of input material, and by using commercial kits; however, these

methods may require specialized equipment not present in every

laboratory (such as refrigerated ultracentrifuges) and can become

increasingly expensive. Another successful and popular approach

for increasing yield is to use strong cell and tissue homogenization

methods such as those based on agitation with microbeads (Fawley

and Fawley, 2004). Automated homogenization has become a stan-

dard step in DNA extraction protocols coupled with second-gen-

eration sequencing platforms, characterized by read sizes under 1

kbp (454 sequencing, Roche, Basel, Switzerland; SOLiD, Illumina,

San Diego, California, USA); however, bead-based homogenization

methods mechanically damage DNA. e resulting low-molecu-

lar-weight DNA is not suitable for third-generation sequencing plat-

forms (Gumińska etal., 2018) unless post-extraction size selection

steps are completed (e.g., dedicated magnetic bead kits or gel-based

systems such as BluePippin [Sage Science, Beverly, Massachusetts,

USA]).

Here, we present a low-cost, highly scalable DNA extraction

protocol specically designed for extracting high-quality, HMW

DNA suitable for use with next-generation long-read sequencing

technologies. Our approach, which we successfully demonstrate in

a variety of green microalgae, optimizes cell lysis to increase yields

while maintaining DNA integrity. First, we compared three meth-

ods for homogenizing and disrupting microalgal cells prior to DNA

extraction, with the aim of maximizing the yield of HMW DNA

without compromising purity. en, we validated the suitability

of our extraction method for application to a broad range of taxa.

We tested the method in a suite of green microalgae within the

Scenedesmaceae (Chlorophyta), which have specialized physiolo-

gies resulting from adaptation to the drastically dierent habitats

of freshwater environments and desert soils. Finally, we veried the

scalability of the method by evaluating the eect of increasing the

initial material input on quality parameters.

METHODS AND RESULTS

Microalgal strains

• Enallax costatus (Schmidle) Pascher, 1943 (isolate CCAP276-31

from the Culture Collection of Algae and Protozoa)

• Tetradesmus obliquus (Turpin) M. J. Wynne, 2016 (isolate Utex

72 from the University of Texas Culture Collection)

• Acutodesmus deserticola (L. A. Lewis & Flechtner ex E. Hegewald,

C. Bock & Krienitz) E. Hegewald, C. Bock & Krienitz, 2013 (iso-

late BCP-SNI-2 from L. Lewis, University of Connecticut)

• Flechtneria rotunda Sciuto & L. A. Lewis, 2015 (isolate BCP-

SEV3-VF49 from L. Lewis, University of Connecticut)

Culturing techniques

Two aquatic (E. costatus and T. obliquus) and two terrestrial (A. de-

serticola and F. rotunda) microalgal species were cultured in 150 mL

of growth medium composed of a 1:1 mix of Bold’s Basal Medium

with micronutrients (Bold, 1949) and Woods Hole Medium (Stein

etal., 1973). All algal cultures were non-axenic monoisolates. All cul-

turing procedures were carried out under sterile conditions. e cul-

tures were grown in 250-mL Erlenmeyer asks at 25°C in a Conviron

PGW36DE growth chamber (Conviron, Winnipeg, Canada) under a

12-h/12-h light/dark photoperiod and 40 μE light from metal halide

and sodium lamps. e cultures were constantly bubbled with am-

bient air. Fresh medium was added every week by allowing the cells

to settle and replacing half of the supernatant (~75 mL) with fresh

medium to sustain high rates of cellular division (Fig.1A). e algal

cultures were grown for six weeks before the DNA extractions.

Cell collection and culture preconditioning

For each algal species and ask, we harvested the cells from the

150-mL culture. We adjusted cultures to a density of ~10

cells

−1

(determined using a Biotek Synergy HT plate reader; BioTek

Instruments, Winooski, Vermont, USA). Algal cells were allowed to

settle and the clear supernatant was poured o. e concentrated al-

gal culture was transferred into a 15-mL Falcon tube, where the cells

were further concentrated by gravity into a nal volume of approxi-

mately 2–3 mL. e remaining supernatant was removed, and 500μL

of each highly concentrated culture were transferred into Eppendorf

tubes for preconditioning prior to the DNA extraction. e samples

were centrifuged for 1 min at 5000 rpm, resulting in the formation of

an algal pellet ranging in size (estimated as volume) from 50–100 μL.

A white layer of debris was observed between the algal pellet and

the supernatant. e composition of this layer was determined un-

der a microscope to be bacteria and empty cell walls (Fig.1A), which

accumulate during cellular division. ese algal species within the

Scenedesmaceae divide asexually through multiple ssion (Cardon

etal., 2018). During this process, a mother cell undergoes multi-

ple rounds of nuclear division followed by cellular division. Once

division is completed, the daughter cells are released, leaving the

empty cell wall of the mother cell behind (Fig.1A). To precondition

Applications in Plant Sciences 2020 8(3): e11333 Stark etal.—High-molecular-weight DNA extraction in microalgae • 3 of 7

the cultures prior to DNA extraction, the supernatant and this de-

bris layer were removed with a micropipette without disturbing the

pelleted cells. e algal pellets were resuspended in 1 mL of fresh

sterile medium by gently inverting the tubes. is preconditioning

process was repeated two additional times to remove additional

debris with varying centrifugation speeds (2500 rpm and again at

5000 rpm; see Appendix1 for step-by-step protocol).

Cell homogenization

For each algal species, we tested the eects of three commonly used

homogenization methods on the quality, molecular weight, and

quantity of the extracted DNA. Success in the homogenization of

cells was initially estimated by visual inspection of the treated sam-

ples under the microscope and by the presence of algal pigments

FIGURE 1. Green microalgal species used in this study and the eect of homogenization methods on the purity of their extracted DNA. (A) Laser scanning

confocal microscope images of the four microalgal species within the Scenedesmaceae used in this study. Images were captured with a 100× objective in

channel mode. The residual light was integrated to generate an optical image. The nuclei were visualized using the dsDNA stain SYBR safe (yellow, 450/50

band pass lter) and the chloroplasts’ chlorophyll uorescence (red, 595/50 band pass lter) (see Cardon etal., 2018 for details). Scale bars = 5 μm. Debris,

cell walls (open arrows), and bacteria (closed arrows) are indicated. Multinucleated cells are dividing cells. (B) Spectral patterns representing absorbance of

a given sample at dierent wave lengths (one representative NanoDrop graph per species per treatment). (C) Scatterplots representing DNA quality mea-

sured as 260 nm/280 nm absorbance ratios vs. the DNA concentration. Grinding with a mini-pestle (squares) or using an automatic vortex adapter (circles)

produced some high-purity, high-concentration samples, but overall the absorbance ratios were lower and more variable. Samples ground in liquid nitro-

gen (LN

; lled diamonds) had uniformly high absorbance ratios across species, even when the pellet sizes were big enough to produce >550 ng/μL DNA.

Applications in Plant Sciences 2020 8(3): e11333 Stark etal.—High-molecular-weight DNA extraction in microalgae • 4 of 7

(e.g., chlorophylls) in the extraction buer. e three homogeniza-

tion methods tested were:

Manual grinding with mini-pestles (two samples per spe-

cies)

—One scoop (~75 μL volume) of autoclaved commercial

silica sand (40–100 mesh, ACROS Organics; ermo Fisher

Scientic, Waltham, Massachusetts, USA) and 100 μL of CTAB

extraction buer (CEB-500-02; OPS Diagnostics, Lebanon, New

Jersey, USA) were added to each algal pellet in an Eppendorf

tube. Each sample was manually ground with a mini-pestle for

approximately 1 min. Resuspending the pellets in small volumes

is required during manual grinding because the cells will oat,

and therefore resuspending in large volumes would decrease the

eectiveness of the process. Aer grinding, 500 μL of CTAB sup-

plemented with 2.5% β-mercaptoethanol (BP176-100; ermo

Fisher Scientic) was added to the cells, for a nal concentration

of 2% β-mercaptoethanol.

Automatic grinding using a vortex adapter (four samples per spe-

cies)

—One scoop of autoclaved commercial silica sand and 600

μL of CTAB extraction buer supplemented with 2.5% β-mercap-

toethanol were added to each algal pellet in an Eppendorf tube.

All samples were placed in a vortex adapter (MoBio 13000-V1-24;

QIAGEN, Hilden, Germany) and vortexed at maximum speed for

5 min. To increase cell lysis, the samples were incubated in a heat

block at 55–60°C for 20 min, followed by an additional 5 min of

vortexing to ensure that enough cells were disrupted.

Grinding in liquid nitrogen (10 samples per species)—Aer pre-

conditioning, all the algal pellets from a single ask (see precon-

ditioning step) were transferred into a pre-chilled mortar using

100–1000-μL wide-bore pipette tips (tip ends were cut with a

sterile blade). If needed, a small volume of the algal medium was

added to the pellets to facilitate the transfer (<100 μL). Samples

were ash-frozen in liquid nitrogen (LN

), and each pellet was

continuously ground using a mini-pestle until the LN

had evap-

orated but the sample had not thawed. e grinding process was

repeated a total of six times, aer which all algal material had

the consistency of ne powder. e number of repetitions was

initially determined by observing the integrity of the cells under

a microscope, and it might vary for other algal taxa. A wide-bore

pipette tip was used to transfer the homogenized algal material

into a sterile Eppendorf tube and centrifuged briey to collect the

sample. No supernatant was removed at this time. e nal vol-

ume of material in each Eppendorf tube was 50–100 μL. Finally,

the samples were frozen in LN

and thawed at room temperature

ve times to lyse any remaining cells. e pellets were gently re-

suspended in 600 μL of CTAB extraction buer supplemented

with 2.5% β-mercaptoethanol. See Appendix1 for a detailed out-

line of this method.

DNA extraction and testing

For the DNA extraction of all samples, regardless of homogeni-

zation method, a modied CTAB extraction method (Doyle and

Doyle, 1987) was used; see Appendix1 for details. roughout the

extraction protocol, we recommend taking general precautions to

prevent additional DNA fragmentation such as minimizing vortex-

ing, gently pipetting (using wide-bore tips if possible), and avoiding

freeze-thaw cycles on extracted DNA.

e homogenized algal cells were incubated in extraction buf-

fer (CTAB supplemented with 2.5% β-mercaptoethanol) at 55–

60°C in a thermal block for 1 h. e tubes were allowed to cool

and reach room temperature; 700 μL of 25:24:1 phenol:chlo-

roform:isoamyl alcohol (IB05174; IBI Scientic, Dubuque, Iowa,

USA) was then added to each tube and briey vortexed to mix.

e samples were centrifuged at 14,000 rpm for 10 min, and the

aqueous upper phase was transferred to a new Eppendorf tube; 4

μL of RNase A (10 mg/mL) (Zymo Research, Irvine, California,

USA) was then added to each sample. e samples were incu-

bated at 37°C for 30 min in a thermal block. e samples were

again allowed to cool to room temperature before a second wash

with 700 μL of phenol: chloroform:isoamyl alcohol and cen-

trifuged. If the samples appeared visibly dirty, this cleaning step

was repeated a third time. e remaining upper aqueous phase

(~350–450 μL) was transferred to a new Eppendorf tube, and the

DNA was precipitated with ~0.1 volumes of 3 M sodium acetate

and ~0.7 volumes of cold isopropanol (Sigma-Aldrich, St. Louis,

Missouri, USA). e samples were mixed by gently inverting the

tubes. Immediately aer mixing, the DNA was easily observable

in most samples as clear, gelatinous blobs at the bottom of the

tube. e samples were stored at −20°C overnight.

Following precipitation, the DNA was pelleted by centrifug-

ing for 5 min at 14,000 rpm. e supernatant was removed, and

the DNA pellets were washed twice with 700 μL of 70% ethanol

(ermo Fisher Scientic). Aer the nal centrifugation step, the

supernatants were removed, and pellets were air-dried by inverting

the tubes on a clean paper towel. e DNA pellets were then resus-

pended in 45 μL of TE buer (ermo Fisher Scientic).

In an additional experiment focused on increasing yield, DNA

was extracted from E. costatus and A. deserticola cultures using the

homogenization method with double the amount of starting

material (“high input”; pellets of ~200 μL). For these high-input

samples, the cells used had been previously frozen at −80°C in cul-

turing medium immediately aer collection. All other steps were

performed without change.

Yield, purity, and integrity of extracted DNA

Yield

—e DNA concentration of each sample was determined

using a Qubit dsDNA HS assay kit (ermo Fisher Scientic)

in a Qubit 2.0 uorometer (ermo Fisher Scientic). Grinding

with LN

produced the highest average DNA yields of the three

methods in T. obliquus (412 ng/μL) and A. deserticola (228 ng/

μL) (Table1). In F. rotunda and E. costatus, the automatic grind-

ing produced the most DNA (272 and 121 ng/μL, respectively),

followed by grinding with LN

(222 and 77 ng/μL, respectively)

(Table1). Grinding samples with mini-pestles resulted in the low-

est DNA concentrations in all species (12–76 ng/μL) (Table1).

e high-input samples (with approximately double the start-

ing algal material) produced higher yields; this was especially

substantial in A. deserticola, with a two-fold change in yield per

Eppendorf tube (Table1).

Purity—DNA absorbance ratios (260 nm/280 nm and 260 nm/230

nm) were determined for all samples using a NanoDrop 2000 mi-

crovolume spectrophotometer (ermo Fisher Scientic) (Fig.1B).

As a general rule, values under 1.8 of the 260 nm/280 nm ratio

indicate carry-over contamination from protein, carbohydrates,

polyphenols, or phenol that could strongly aect downstream

Applications in Plant Sciences 2020 8(3): e11333 Stark etal.—High-molecular-weight DNA extraction in microalgae • 5 of 7

applications. Values of ~1.8 are considered pure DNA, whereas

values over 1.8 can be associated with changes in nucleotide ratios,

especially increases in adenine content (chloroplast genomes, for

example, are adenine rich [Smith, 2012]), or the presence of RNA.

If present, RNA can be observed easily in agarose gels and removed

by means of additional RNase A treatment of samples followed by

ethanol precipitation. We did not observe presence of RNA when

our samples were checked on agarose gels (see Fig. 2A, C for pos-

itive and negative images of representative gels), so no additional

RNase A treatments were performed. For the 260 nm/230 nm ra-

tio, values under 2.0 indicate presence of contaminants. For all the

treatments, the 260 nm/280 nm ratios ranged between 1.8 and 2.1.

e absorbance ratios of those DNAs extracted by grinding with

mini-pestles were consistently lower (1.8–1.9) than those extracted

using automatic grinding or LN

(1.9–2.1) (Table1, Fig.1C). For all

species, the 260 nm/230 nm ratio was over 2 in samples homoge-

nized with LN

, as is recommended for long-read sequencing. In E.

costatus and F. rotunda, lower 260 nm/230 nm ratios were observed

for samples extracted using other lysis methods.

e absorbance ratios were compared across treatments using

an ANOVA performed in R (R Development Core Team, 2011)

(Table 1). When pooling all species, dierent homogenization

methods produced signicantly dierent ratios of absorbance at

260 nm/230 nm (F

2,70

= 26.88, P < 0.001) and at 260 nm/280 nm

2,70

= 165.8, P < 0.001). LN

grinding produced the highest 260

nm/230 nm ratios for all species (1.9–2.4; see Table1), including in

those samples in which the initial amount of algal material had been

doubled (Table1).

Integrity—e molecular weight of the extracted DNA was deter-

mined by running 2 μL of each sample in 0.8% agarose gel, 1× TAE

buer (ermo Fisher Scientic) for 4 h at 60 V. e DNA was stained

with 1× SYBR safe (Invitrogen, Waltham, Massachusetts, USA). e

molecular weight of the genomic DNA was estimated by compar-

ison with a HMW DNA ladder (Lambda DNA/EcoRI+HindIII;

ermo Fisher Scientic). Digital images of the gels were gener-

ated in an Amersham 600 RGB imager (GE Healthcare, Chicago,

Illinois, USA) using automatic collection parameters. e images of

the gels were enhanced (contrast, homogenization, and background

removal) in ImageJ version 1.52a (Rasband, 2018) prior to the dig-

ital analysis. e 1D gel electrophoresis image analysis soware

GelAnalyzer2010a (www.gelan alyzer.com) was used to create pro-

les of the distribution of DNA fragments (Fig.2A, B) using the

Lambda DNA/EcoRI+HindIII DNA ladder as a size reference.

For all species, HMW DNA was exclusively observed in the DNA

samples extracted from cells homogenized using the LN

grinding

method. e DNA extracted using this treatment was observed as

a tight, clear band over the 21.2-kbp marker band, whereas DNA

extracted from cells homogenized using the other treatments dis-

played substantial smearing and lacked a clear HMW DNA band,

consistent with high DNA fragmentation (Fig.2).

CONCLUSIONS

Current genome sequencing protocols rely on a combined ap-

proach of short (Illumina) and long (PacBio, Nanopore) reads. e

eciency of long-read sequencing is directly impacted by the integ-

rity of the DNA used. Our results indicate that for all algal species

tested here, a modied CTAB protocol is sucient for extracting

DNA within reasonable quality parameters. However, DNA integ-

rity is strongly aected by the cellular homogenization method used

early in the extraction protocols (i.e., during the lysis step). DNA ex-

tracted using automatic or mini-pestle grinding is suitable for PCR

or short-read sequencing but not for long-read sequencing technol-

ogies. Grinding cells in LN

was the only homogenization method

that consistently resulted in HMW DNA.

Many potential modications to DNA extraction methods

and kits can produce acceptable results for dierent sequencing

methods; however, they can be much more expensive and less

customizable than CTAB-based extraction protocols. Our results

demonstrate that, for a diverse suite of microalgae taxa that in-

cludes aquatic species and desert-derived species with recalcitrant

characteristics for DNA extraction, sample preparation and cell ly-

sis methods were key to producing high-quality DNA. Across the

four species, uniformly good results were obtained from the CTAB

extraction aer grinding the cells in LN

, even though the initial

samples varied in cell size, cell wall thickness, and buoyancy. We also

TABLE 1. Quality parameters measured for DNA extracted from cells homogenized using dierent methods.

  Concentration (Qubit) Purity (NanoDrop)

Integrity

(electrophoresis)

Species (habitat) Method (n) Mean (ng/μL ± SE) Total (μg)

260 nm/280 nm

(mean ± SE)

260 nm/230 nm

(mean ± SE)

Distribution of DNA

fragments

Enallax costatus

(aquatic)

Pestle (2) 49 ± 4 2.2 1.9 ± 0.02 1.6 ± 0.11 Broad peak

Auto (4) 121 ± 13 5.4 2.0 ± 0.01 1.2 ± 0.03 Broad peak

(10) 77 ± 11 3.4 2.1 ± 0.01 2.2 ± 0.03 Tight peak

High input (5) 103 ± 9 4.6 2.1 ± <0.01 1.9 ± 0.02 Tight peak

Tetradesmus

obliquus

(aquatic)

Pestle (2) 12 ± 2 0.5 1.9 ± 0.02 2.1 ± 0.05 Broad peak

Auto (4) 198 ± 55 8.9 2.0 ± 0.01 2.1 ± 0.04 Broad peak

(6) 412 ± 27 18.5 2.1 ± 0.01 2.4 ± 0.01 Tight peak

Acutodesmus

deserticola

(desert)

Pestle (2) 35 ± 1 1.5 1.9 ± <0.01 2.1 ± 0.03 Broad peak

Auto (4) 66 ± 13 2.9 2.0 ± 0.01 2.0 ± 0.04 Broad peak

(10) 228 ± 27 10.2 2.1 ± <0.01 2.3 ± 0.01 Tight peak

High input (6) 448 ± 41 20.1 2.1 ± <0.01 2.3 ± 0.01 Tight peak

Flechtneria

rotunda (desert)

Pestle (2) 76 ± 10 3.4 1.8 ± <0.01 1.3 ± 0.03 Broad peak

Auto (4) 272 ± 46 12.2 1.9 ± 0.01 1.8 ± 0.05 Broad peak

(12) 222 ± 42 9.9 2.1 ± 0.01 2.2 ± 0.03 Tight peak

Note: n = number of samples.

For each sample, the total volume was 45 μL.

Applications in Plant Sciences 2020 8(3): e11333 Stark etal.—High-molecular-weight DNA extraction in microalgae • 6 of 7

demonstrated that this method can produce up to 20 μg of DNA in

a single Eppendorf tube without sacricing purity or quality, using

fresh or frozen material.

ACKNOWLEDGMENTS

e authors thank Dr. Louise Lewis (University of Connecticut) for

providing Flechtneria rotunda and Acutodesmus deserticola, and

Suzanne omas for expert technical assistance. is work was sup-

ported by the National Science Foundation, Division of Integrative

Organismal Systems (1355085 to Z.G.C.) and an anonymous donor

(to Z.G.C.).

AUTHOR CONTRIBUTIONS

E.L.P. and Z.G.C. designed the experiments; E.L.P. and J.R.S. ac-

quired and analyzed the data; E.L.P. interpreted the data; J.R.S. and

E.L.P. wrote the manuscript; and J.R.S., Z.G.C., and E.L.P. approved

the submitted version.

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APPENDIX 1. DNA extraction protocol: Sample homogenization using liquid

nitrogen and modied CTAB method.

1. Preconditioning of cultures

a. Concentrate algal cultures in the minimum volume

possible and aliquot (500 μL) into Eppendorf tubes

for preconditioning

b. Centrifuge algal cells in growth medium at 5000 rpm

for 1 min

c. Carefully remove the supernatant and white interface

layer (cell walls and bacteria) with a micropipette

d. Add 1 mL of fresh sterile growth medium and resus-

pend cells

e. Centrifuge at 2500 rpm for 30 s

f. Repeat steps c–d

g. Centrifuge at 5000 rpm for 30 s

h. Repeat step c, removing as much supernatant as possible

2. Cell grinding with liquid nitrogen (LN

)

a. Autoclave mortar and pestle to sterilize

b. Pre-chill mortar and pestle with LN

c. Resuspend algal culture in as little medium as possible

(here <100 μL) and transfer to pre-chilled mortar using

a wide-bore tip (cut tip end with sharp, sterile blade)

d. Grind with pestle until LN

has evaporated but cells

have not thawed

e. Add a small amount of additional LN

f. Repeat steps d and e ve more times (cells should

look damaged under microscope)

g. Transfer algal material to Eppendorf tubes and cen-

trifuge briey to collect sample in bottom

h. Freeze and thaw (at room temperature) centrifuge

tubes ve times in LN

3. DNA extraction

a. Prepare extraction buer with CTAB and 2.5% β-mer-

captoethanol. Add enough CTAB- β-mercaptoethanol

to each Eppendorf tube to bring the total volume to

600 μL

b. Incubate samples at 55–60°C

c. Add 700 μL of phenol:chloroform:isoamyl alcohol

(25:24:1) and vortex for 3–5 s to mix

d. Centrifuge for 10 min at 14,000 rpm

e. Transfer aqueous phase to new sterile Eppendorf and

add 4 μL of RNase A

f. Incubate for 30 min at 37°C

g. Repeat steps b–d and transfer aqueous phase to new sterile

Eppendorf; repeat twice if samples still appear visibly dirty

h. Add 0.1 volumes of cold 3 M sodium acetate and 0.7–

0.9 volumes of cold isopropanol. Mix by inversion.

i. Precipitate overnight at −20°C

j. Centrifuge for 5 min at 14,000 rpm

k. Remove supernatant

l. Add 700 μL of cold 70% ethanol to wash DNA pellet

m. Centrifuge for 1 min at 14,000 rpm

n. Repeat steps k–m

o. Remove supernatant and invert Eppendorf tubes on a

clean paper towel to air-dry pellets

p. Resuspend in 45 μL of TE buer in refrigerator at

least overnight