Reproduction, hatchery and culture applications for the giant kelp (Macrocystis pyrifera): a methodological appraisal Reproduction, hatchery and culture applications for the giant kelp (Macrocystis pyrifera): a methodological appraisal

Abstract

2022) Reproduction, hatchery and culture applications for the giant kelp (Macrocystispyrifera): a methodological appraisal, Applied Phycology, 3:1, 368-382, ABSTRACT Although much is known regarding the physiology, ecology and life history of Macrocystis pyrifera, there is little accessible information for establishing robust and reliable culturing practices to support aquaculture and habitat restoration. Naturally occurring kelp forests formed by M. pyrifera support productive coastal ecosystems, and because it is one of the fastest growing macroalgal species, its high biomass production, and high affinity for nutrients and significant polysaccharide content makes it a species of considerable interest for aquaculture. This species has undergone substantial decline throughout its biogeographic range and is threatened by local and global stressors. Here, we synthesize the current knowledge on culturing of M. pyrifera and discuss approaches to stock collection, preservation of diversity and applications for experimental studies. It is crucial to preserve the current genetic diversity of this species immediately and long-term culture storage approaches such as germplasm banking and cryopreservation provide the tools to allow this. A concerted effort is also needed to better understand the physiological attributes of M. pyrifera in order to select strains for aquaculture and restorative applications that may provide resilience to future environmental stressors. Finally, attention must be given to developing effective in situ restoration approaches whereby large-scale stock production can be optimized and out-planting strategies developed to ensure restoration success. ARTICLE HISTORY

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Reproduction, hatchery and culture applications
for the giant kelp (Macrocystis pyrifera): a
methodological appraisal
Duong M. Le, Matthew J. Desmond, Alejandro H. Buschmann, Daniel W.
Pritchard, Carolina Camus, Catriona L. Hurd & Christopher D. Hepburn
To cite this article: Duong M. Le, Matthew J. Desmond, Alejandro H. Buschmann, Daniel W.
Pritchard, Carolina Camus, Catriona L. Hurd & Christopher D. Hepburn (2022) Reproduction,
hatchery and culture applications for the giant kelp (Macrocystis�pyrifera): a methodological
appraisal, Applied Phycology, 3:1, 368-382, DOI: 10.1080/26388081.2022.2086823
To link to this article: https://doi.org/10.1080/26388081.2022.2086823
© 2022 The Author(s). Published by Informa
UK Limited, trading as Taylor & Francis
Group.
Published online: 04 Sep 2022.
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Reproduction, hatchery and culture applications for the giant kelp (Macrocystis
pyrifera): a methodological appraisal
Duong M. Le
a,b
, Matthew J. Desmond
a,b
, Alejandro H. Buschmann
c
, Daniel W. Pritchard
a
, Carolina Camus
c
,
Catriona L. Hurd
d
and Christopher D. Hepburn
a,b
a
Department of Marine Science, University of Otago, Dunedin, New Zealand;
b
Coastal People Southern Skies Centre of Research Excellence,
University of Otago, Dunedin, New Zealand;
c
Centro I~mar and CeBiB, Universidad de Los Lagos, Puerto Montt, Chile;
d
Institute for Marine and
Antarctic Studies, University of Tasmania, Hobart, Tasmania, Australia
ABSTRACT
Although much is known regarding the physiology, ecology and life history of Macrocystis pyrifera,
there is little accessible information for establishing robust and reliable culturing practices to
support aquaculture and habitat restoration. Naturally occurring kelp forests formed by M. pyrifera
support productive coastal ecosystems, and because it is one of the fastest growing macroalgal
species, its high biomass production, and high anity for nutrients and signicant polysaccharide
content makes it a species of considerable interest for aquaculture. This species has undergone
substantial decline throughout its biogeographic range and is threatened by local and global
stressors. Here, we synthesize the current knowledge on culturing of M. pyrifera and discuss
approaches to stock collection, preservation of diversity and applications for experimental studies.
It is crucial to preserve the current genetic diversity of this species immediately and long-term
culture storage approaches such as germplasm banking and cryopreservation provide the tools to
allow this. A concerted eort is also needed to better understand the physiological attributes of
M. pyrifera in order to select strains for aquaculture and restorative applications that may provide
resilience to future environmental stressors. Finally, attention must be given to developing
eective in situ restoration approaches whereby large-scale stock production can be optimized
and out-planting strategies developed to ensure restoration success.
ARTICLE HISTORY
Received 26 December 2021
Accepted 30 May 2022
KEYWORDS
Aquaculture; giant kelp;
hatchery; long-term storage;
Macrocystis pyrifera;
reproduction; restoration
Introduction
Farming of marine macroalgae has a long history
but has seen a significant increase in scale and
advancement in research and infrastructure since
the middle of the twentieth century. In 2018, the
global production of macroalgae reached more than
32.4 million tonnes per annum (FAO, 2020a), with
97.1% coming from farmed systems (Chopin &
Tacon, 2021). 45.3% of that production came from
cultivated kelp species of the order Laminariales
(FAO, 2020a). The main focus of macroalgal pro-
duction is on polysaccharide products followed by
direct human consumption (Naylor et al., 2021).
However, in recent years, and in response to envir-
onmental challenges, there has been a shift in
research focus to better understand culturing prac-
tices for other uses, such as multitrophic aquacul-
ture, preserving and buffering ecosystem function,
maintaining biodiversity and restoration (Layton
et al., 2020; Ling, Johnson, Frusher, & Ridgway,
2009; Terawaki, Hasegawa, Arai, & Ohno, 2001).
Of all macroalgae, kelps (Laminariales) are gen-
erally the largest and exhibit some of the highest
productive yields (Bolton, Anderson, Smit, &
Rothman, 2012; Reed, Rassweiler, & Arkema,
2008). They are found throughout temperate
regions and provide key services to nearshore and
coastal systems through their role as ecosystem
engineers (Bolton et al., 2012; Graham, Vásquez, &
Buschmann, 2007). Globally, 38% of ecoregions
have exhibited kelp forest decline over the past
50 years (Krumhansl et al., 2016). Multiple drivers,
such as increased ocean temperature (Filbee-Dexter,
Feehan, & Scheibling, 2016; Johnson et al., 2011;
Wernberg et al., 2013), sedimentation (Connell,
2003; Filbee-Dexter & Wernberg, 2018), eutrophica-
tion (Filbee-Dexter & Wernberg, 2018), overgrazing
(Kriegisch, Reeves, Johnson, & Ling, 2019; Ling
et al., 2009), invasion (Blamey & Branch, 2012)
and overfishing (Andrew & O’Neill, 2000; Ling
et al., 2009; Mabin, Johnson, & Wright, 2019) are
to blame for continued decline.
CONTACT Duong M. Le ledu7263@student.otago.ac.nz
APPLIED PHYCOLOGY
2022, VOL. 3, NO. 1, 368–382
https://doi.org/10.1080/26388081.2022.2086823
British
Phycological
Society
Understanding and using algae
© 2022 The Author(s). Published by Informa UK Limited, trading as Taylor & Francis Group.
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use,
distribution, and reproduction in any medium, provided the original work is properly cited.

The giant kelp, Macrocystis pyrifera, forms and main-
tains the largest ecosystems of all macroalgae, creating
forests that modify the physical and chemical environ-
ment, providing key habitat, food resources and services
to a plethora of species (Graham et al., 2007). M. pyrifera
holds significant value as a direct harvest resource, pre-
dominantly for the extraction of the polysaccharide
alginate (McHugh, 2003; Ortiz et al., 2009), abalone
feed (Correa et al., 2016), as an additive for liquid
fertilizer production (Gutierrez et al., 2006) and for its
role in supporting cultural, recreational and commer-
cially important fisheries (Jones, Lawron, & Shachak,
1997; Wernberg, Krumhansl, Filbee-Dexter, &
Pedersen, 2019). Historically, the majority of harvests
have been taken from wild populations (Buschmann
et al., 2004; Purcell-Meyerink, Packer, Wheeler, &
Hayes, 2021), with Chile and Peru being the main pro-
ducers (i.e., 33 979 and 32 794 tonnes, respectively, in
2019; Cai et al.,). Like many other kelp species, wild
M. pyrifera forests have undergone significant global
decline (Hay, 1990; Johnson et al., 2011; Krumhansl
et al., 2016; Filbee-Dexter & Wernberg, 2018; Tait
et al., 2021) and as a result, many of the services they
provide have declined or been lost.
As attention shifts more towards the culturing of
M. pyrifera for both harvest and bioremediation and
restoration purposes, it has become clear that there is
a lack of comprehensive information regarding cultur-
ing practices for this important species. Many studies
have investigated the life history of M. pyrifera and by
contrasting these bodies of work it is apparent that the
conditions needed for successful growth are variable
from one region to another (Gutierrez et al., 2006;
Leal, Roleda, Fernández, Nitschke, & Hurd, 2021;
Lüning, 1981; Macchiavello, Araya, & Bulboa, 2010;
Neushul, 1963; Schiel & Foster, 2015; Westermeier,
Patiño, Piel, Maier, & Mueller, 2006), making it difficult
to establish robust and reliable culturing practices.
Approaches to culturing a particular species are also
highly dependent on the purpose of culturing. This
critical assessment aims to synthesize the current meth-
ods and conditions used when culturing M. pyrifera,
with a specific focus on the needs and application of
the cultured product. This assessment will discuss meth-
ods for stock collection, storage and grow-out, with
reference to experimental, aquacultural and restorative
applications.
Life history and distribution
M. pyrifera is found in temperate waters along the coasts
of Southern Africa, east and west South America,
Tasmania and South Australia, central and southern
New Zealand (including the sub Antarctic Islands) and
the west coast of North America and Canada (Figure 1).
It requires rocky substrata for attachment and can be
found in both sheltered and exposed environments
(Graham et al., 2007; Schiel & Foster, 2015).
Figure 1. Global distribution of Macrocystis pyrifera (modified from Graham et al., 2007).
APPLIED PHYCOLOGY 369

Reproduction of M. pyrifera can occur year round
(Brown, Nyman, Keogh, & Chin, 1997; Reed, Ebeling,
Anderson, & Anghera, 1996) but the abundance of
reproductive tissue (sorus) and density of zoospores
(hereafter spores) within sori varies seasonally
(Buschmann et al., 2004; Leal et al., 2021; Neushul,
1963; Reed et al., 1996). Like other Laminariales,
M. pyrifera has a biphasic life cycle, alternating between
a microscopic haploid stage and a macroscopic diploid
stage (Schiel & Foster, 2015). The main reproductive
tissue is located on the sporophyll at the base of the
sporophyte, above the holdfast, although spore bearing
tissue can also be found in the blades and the apical
scimitar of some individuals (Leal, Hurd, & Roleda,
2014; Leal et al., 2021; Neushul, 1963).
In the wild, spore reproduction occurs throughout
the year, with the highest production coinciding with
low-temperature months, such as early winter and late
spring/early summer (Anderson & North, 1967; Reed
et al., 1996), and storms (Reed et al., 2006). Mature sori
release spores that swim freely in the water column and
are transported by currents with a typical dispersal
range of <1 km. Spore settlement occurs within hours
to a few days post release (Devinny & Volse, 1978; Reed,
Amsler, & Ebeling, 1992) and is triggered by the avail-
able nutrients (i.e., by following a concentration gradi-
ent) (Amsler & Neushul, 1989), presence of other biota
(Reed et al., 1996), and light condition (Reed et al.,
1992). Germination can occur immediately after spore
release and is typically complete within 48 hrs under
laboratory conditions (Anderson & Hunt, 1988;
Garman, Pillai, Goff, & Cherr, 1994), or a few days
post release in the field (Devinny & Volse, 1978; Reed
et al., 1992; Santelices, 1990). After germination, germl-
ings start to grow by increasing cell number and size,
forming male and female gametophytes (Devinny &
Volse, 1978; Reed et al., 1992; Schiel & Foster, 2015).
In optimal laboratory conditions, gametophytes of both
sexes can become fertile in approximately two weeks
(Lüning & Neushul, 1978). The fertilization process
takes place when eggs of female gametophytes produce
a pheromone called lamoxirene, which attracts the
sperm of male gametophytes (Maier, Hertweck, &
Boland, 2001). The effective distance for hormone
attraction is reported to be within 1 mm (Boland,
Marner, Jaenicke, Muller, & Folster, 1983). If successful
fertilization occurs, then female gametophytes turn into
embryonic sporophytes.
Once established, the growth of sporophytes can be
rapid. Sporophyte growth can result in a doubling of the
length every month from an initial length of 1–2 cm
(Neushul, 1963). Growth rates are variable across season
and by region (North, 1976a; Wheeler & North, 1981;
González-Fragoso, Ibarra-Obando, & North, 1991;
Hernández-Carmona, 1996; Brown et al., 1997;
Graham et al., 2007; Macchiavello et al., 2010), for
example, greatest growth rates occur during the summer
in Alaska and British Columbia, while populations in
California and New Zealand displayed greatest growth
during the winter and spring (North, 1976a; Wheeler &
North, 1981; Brown et al., 1997; Graham et al., 2007).
This variation in growth is driven by resource
Figure 2. Lifecycle of Macrocystis pyrifera.
370 D. M. LE ET AL.

availability, namely, light and nutrients (Desmond,
Pritchard, & Hepburn, 2015; Graham et al., 2007;
Schiel & Foster, 2015; Stephens & Hepburn, 2014).
M. pyrifera populations have a relatively high turnover
rate due to their large size, which makes them vulner-
able to physical forces, but individuals have been shown
to persist for up to 7 years (Graham et al., 2007).
Stock collection
In the following section stock refers to the culture of
M. pyrifera from a known source that may be used for
multiple applications as outlined below.
Seasonality and timing
M. pyrifera can be reproductive year-round, but there
appear to be seasonal trends when spore production is
highest, and depending on location, there may be peri-
ods where no sorus tissue is present (Brown et al., 1997;
Buschmann et al., 2004; Hay, 1990; Leal et al., 2021;
Neushul, 1963; Reed et al., 1996). To ensure the greatest
chance of successful spore release, collection should be
carried out when spore production is highest. Highest
sporophyll counts and spore densities are typically
observed during the austral summer and autumn period
in the Southern Hemisphere (Buschmann, Moreno,
Vásquez, & Hernández-González, 2006; Hay, 1990;
Henríquez et al., 2011), while populations along the
west coast of the United States show highest spore
densities during the winter and spring (Reed et al.,
1996).
Collection and transportation
Mature sorus tissue can be removed by cutting it
away from the sporophyll without removing the
entire individual. If possible, immediate sporulation
in the field is the best method to obtain the highest
spore density (Suebsanguan, Strain, Morris, &
Swearer, 2021) but often this is not practical. If it
is not possible to sporulate immediately, then spore
tissue can be transported either semi-dry, packed in
damp paper cloth which maintains higher spore
densities when compared to wet transportation
(Bak, Mols-Mortensen, & Gregersen, 2018; Kim,
Yarish, Hwang, Park, & Kim, 2017; Suebsanguan
et al., 2021), or in a plastic zip-lock bag or similar
sealed container filled with seawater and kept in the
dark at ~4°C (Barrento, Camus, Sousa-Pinto, &
Buschmann, 2016; Gutierrez et al., 2006;
Westermeier et al., 2006). Generally, longer trans-
portation time (>24 hrs) leads to lower spore yield
so facilities and equipment should be well prepared
and sterilized to be ready to receive reproductive
material (North, 1976a; Devinny & Leventhal, 1979;
Gutierrez et al., 2006; Kim et al., 2017; Suebsanguan
et al., 2021) .
Sporulation
Pre-treatment
After transporting the fresh mature sorus to the
laboratory or processing facility, it is important to
clean the tissue to reduce the likelihood of culture
contamination. Typically, cleaning is performed by
gently rinsing the sorus tissue with filtered (0.2 µm
pore diameter) and/or sterilized (autoclaved) sea-
water and wiping off any visible fouling. Tissue can
also be rinsed using freshwater, freshwater with
chlorine (5 ml of chlorine l
–1
water) or iodine with-
out any noted negative effects on spore quality
(Alsuwaiyan et al., 2019; Camus & Buschmann,
2017; Plá & Alveal, 2012). After cleaning, the sorus
tissue should undergo a desiccation period to
enhance spore release (Leal et al., 2014). Most studies
store sorus tissue in damp paper cloth for between 1
and 24 hrs in the dark at temperatures between 4°C
and 15°C (Camus & Buschmann, 2017; Gutierrez
et al., 2006; Leal et al., 2014; Plá & Alveal, 2012).
After the desiccation period, sporulation can take
place. It should be noted that sporulation can take
place without a desiccation period; however, this
approach is more time consuming, has a greater
risk of contamination and produces more mucilage
(Neushul, 1963; Sanbonsuga & Neushul, 1978).
Medium
For sporulation, a range of culture media are reported,
the most common being filtered, sterilized and/or nutri-
ent enriched seawater (i.e., Provasoli, Alsuwaiyan et al.,
2019). Artificial seawater is sometimes used; however,
the advantages or disadvantages of this approach are not
clear (Amsler & Neushul, 1989, 1990). A general guide
should be that the medium used should minimize the
chance of infection or contamination and therefore both
filtration and sterilization either using UV or autoclave
are recommended. Very few studies report using ger-
manium oxide or antibiotics to control contamination
at the time of sporulation, and it is advised that if used
this should be precise in concentration and duration to
reduce negative effect on growth (Kawai, Motomura, &
Okuda, 2005; Markham & Hagmeier, 1982; Shea &
Chopin, 2007).
APPLIED PHYCOLOGY 371

Sporulation time, temperature, and light intensity
The desiccated sorus tissue should be submerged in
the medium and left for a period of time (“sporula-
tion time”) to allow release. Of the studies that
report sporulation time, most range between 15 min-
utes to 1 hr (Camus & Buschmann, 2017; Gutierrez
et al., 2006; Leal et al., 2014; Macchiavello et al.,
2010; Neushul, 1963; Plá & Alveal, 2012). Many
studies do not state the temperature used for spor-
ulation, of those that do, most range between 10°C
and 18°C (Alsuwaiyan et al., 2019; Camus &
Buschmann, 2017; Gutierrez et al., 2006; Leal
et al., 2014; Macchiavello et al., 2010; Neushul,
1963; Plá & Alveal, 2012). The lowest temperature
reported was 0°C (James, Stull, & North, 1990) and
the highest 19.8°C (Hollarsmith, Buschmann,
Camus, & Grosholz, 2020). Little information is
available regarding the potential effects of sporula-
tion temperature on developmental parameters, and
this is an area that requires further research given
its potential ecological importance. Light intensity is
variable across studies, ranging from complete dark-
ness to dim irradiance (Alsuwaiyan et al., 2019).
Most studies did not mention irradiance levels dur-
ing sporulation (Alsuwaiyan et al., 2019). Care
must, however, be taken to avoid potential photo
damage during this time (Graham, 1996), and it is
therefore recommended that sporulation be con-
ducted under dim light conditions.
Spore density
The number of spores per unit area of sorus tissue is
variable between populations (Buschmann et al., 2004),
season (Buschmann et al., 2004; Neushul, 1963) and
tissue types (Leal et al., 2014, 2021; Neushul, 1963);
therefore, the number of spores produced during spor-
ulation will also vary. Spore density can be assessed post
sporulation by taking an aliquot of the medium and
quantifying spore density under magnification using
a haemocytometer. Studies report densities in the
range of 40,000–5,000,000 spores ml
–1
(Barrento et al.,
2016; Buschmann et al., 2006, 2004; Camus &
Buschmann, 2017; Cie & Edwards, 2008; Deysher &
Dean, 1984; Leal et al., 2014; Macchiavello et al., 2010;
Muñoz, Hernández-González, Buschmann, Graham, &
Vásquez, 2004; Westermeier et al., 2006). Spore density
can be diluted by adding an additional medium to the
sporulation vessel in order to achieve the desired con-
centration, which will be highly dependent on the appli-
cation of the cultured stock and will be discussed
further below.
Stock management
It is important to have a clear understanding of the
intended application of cultured M. pyrifera as this
will dictate how much tissue should be collected,
how sporulation should be undertaken and most
importantly what happens to stock post sporulation.
In this section, we discuss long-term storage techni-
ques that allow for the preservation of genetic diver-
sity and culture stocks, short-term storage and
culturing methods for experimental applications,
and finally grow-out approaches for aquacultural
and restoration purposes.
Preserving diversity
The need for long-term storage of M. pyrifera is
multifaceted. Given the decline of M. pyrifera for-
ests worldwide, a loss of genetic diversity has more
than likely occurred. Continued pressure on these
systems means that a concerted effort is needed to
preserve current genetic strains for future uses
(Barrento et al., 2016), whatever they may be. In
situ protection measures, such as marine protected
areas, do not offer the insurance needed to preserve
genetic diversity as these areas remain vulnerable to
the effects of global stressors (Williams, 2001). Ex
situ preservation methods are therefore the only
way to safeguard and preserve diversity (Coleman
et al., 2020; Wade et al., 2020). Large-scale ex situ
preservation of terrestrial plants and crop species
has occurred in climate-controlled seed banks
since the early 20th century in order to ensure
food security and preserve biodiversity (Wade
et al., 2020). Long-term storage of macroalgae, how-
ever, lags significantly behind terrestrial examples,
but two main approaches are slowly becoming
utilized.
Germplasm banking
Germplasm banking is defined as the preservation of
biological, animal or plant, for the purposes of
delayed breeding and propagation. For macroalgae,
germplasm banking involves holding male and female
gametophytes, often separately, in a state of dormancy
by placing them under reduced temperature, light and
nutrient conditions (Barrento et al., 2016). For most
kelps, including M. pyrifera, the gametophytic (hap-
loid) stage responds best to long-term storage condi-
tions (Barrento et al., 2016; Carney & Edwards, 2010;
Westermeier et al., 2006). Only a small number of
studies have reported storing M. pyrifera under
372 D. M. LE ET AL.

germplasm conditions (Barrento et al., 2016; Lewis &
Neushul, 1994; Westermeier et al., 2006; Xu et al.,
2015).
The key factors found to affect the survivorship
of M. pyrifera gametophytes under germplasm con-
ditions are temperature, light, nutrient concentra-
tion and cell density. Ideally, germplasm storage
temperature should be at the low end of the natu-
rally occurring temperature range for the region
where stock originates. Temperature conditions
vary between 8°C and 15°C which is considered
very close to the optimal growth range for
M. pyrifera (i.e., 12–17°C, Lüning & Neushul,
1978) and so the temperature for germplasm should
be lower than 12°C. Importantly, light levels must
be kept low, <5 µmol m
–2
s
–1
is most common, and
light:dark photoperiods between 10 L:14D –
16 L:8D are recommended (Barrento et al., 2016;
Westermeier et al., 2006; Xu et al., 2015). The
removal of blue wavelength light from the light
source (e.g., using red light LEDs) provides further
insurance to avoid gametophytic sexual develop-
ment (Lüning & Neushul, 1978).
Nutrient availability is particularly important for
controlling vegetative growth and maintaining germ-
plasm cultures at manageable densities. Cultures
should be held in nutrient enriched, filtered, auto-
claved seawater with the preferred medium being
Provasoli (Barrento et al., 2016; Carney & Edwards,
2010; Wade et al., 2020; Westermeier et al., 2006). If
cultures are maintained at low densities, the medium
can be changed approximately twice per year as long
as evaporation is kept to a minimum (Barrento et al.,
2016; Westermeier et al., 2006). To completely pre-
vent any chance of reproduction, it is advised that
male and female gametophytes are held separately
once distinguishable. The 96-well plates are com-
monly used to store gametophytes as they offer
highly independent storage replication at low densi-
ties. Studies that place M. pyrifera gametophytes in
long-term storage have shown viability of stock after
more than 5 years post preservation (Barrento et al.,
2016; Lewis & Neushul, 1994; Westermeier et al.,
2006; Xu et al., 2015).
Cryopreservation
Cryopreservation is a second approach, which is
gaining attention for its potential as a long-term
storage option for algal cultures (Choi, Nam, &
Kuwano, 2013; Kuwano, Kono, Jo, Shin, & Saga,
2004; Vigneron, Arbault, & Kaas, 1997; Zhang
et al., 2007; Zhang, Cong, Qu, Luo, & Yang, 2008).
Cryopreservation was first used to preserve human
sperm in the 1960ʹs (Sherman, 1973) and has since
been used in both agricultural and horticultural
practices to preserve biological material (Taylor &
Fletcher, 1998). The process, in general, requires the
staged cooling of the subject to very low tempera-
tures where biological cell degradation effectively
ceases (Grout, 1995). Typically, the samples are
stored under solid carbon dioxide or liquid nitrogen
conditions to achieve temperatures of – 80 to –
196°C. As with germplasm banking, the gametophy-
tic stage of M. pyrifera responds best to cryopreser-
vation (Piel, Avila, & Alcapán, 2015).
The process of preservation requires gameto-
phytes to be suspended in a cryoprotectant solution
made up of dimethyl sulphoxide (DMSO), glycerol,
sucrose, dextrose and sorbitol (Piel et al., 2015;
Zhang et al., 2007, 2008) before it is cooled. The
cooling step is separated into two minor steps to
avoid the damage to cell walls (Piel et al., 2015;
Zhang et al., 2007, 2008). Briefly, gametophyte sus-
pensions are pre-frozen either at –18°C to –20°C for
30 min. After 30 min pre-frozen samples are placed
in liquid nitrogen (Piel et al., 2015; Zhang et al.,
2007). Only one published study considers the cryo-
preservation of M. pyrifera (Piel et al., 2015). Four
weeks after being thawed and returned to optimal
growing conditions 29% of female gametophytes
were still immature and in a vegetative state, 9%
released eggs and 9% developed into embryonic
sporophytes. From other species, it appears that
survival rate and viability of preserved tissue is
species dependent (Ginsburger-Vogel, Arbault, &
Pérez, 1992; Kono, Kuwano, & Saga, 1998;
Kuwano et al., 2004; Piel et al., 2015; Zhang et al.,
2007, 2008) and is significantly influenced by the
rate of freezing and thawing process (Zhang et al.,
2007). Therefore, more research is needed to opti-
mize this approach to enable the successful long-
term storage of M. pyrifera.
Both germplasm banking and cryopreservation
allow for the long-term storage of M. pyrifera,
which is essential for preserving the genetic diver-
sity of this species. Government, research agencies
and other parties interested in the values surround-
ing kelp forests should make a concerted effort to
establish long-term storage facilities to preserve this
species. Not only does storage safeguard genetic
diversity but it also forms the basis for breeding
programs and allows for timely propagation of
stock for experimental, aquacultural and restorative
applications.
APPLIED PHYCOLOGY 373

Farming and applications
Culturing methods for laboratory experimentation
Laboratory experimentation is important to understand
the physiological function of M. pyrifera and knowledge
gained often provides perspective on ecological trends
or guide approaches for other applications, such as
aquaculture or restoration. Typically, these types of
experiments focus on the early life stages from the
spore through to the juvenile sporophyte (Barrento
et al., 2016; Carney & Edwards, 2010; Hollarsmith
et al., 2020; Leal, Hurd, Fernández, & Roleda, 2017;
Westermeier et al., 2006). This tends to be because of
the importance of these early life stages in future devel-
opment (Schiel & Foster, 2015) as well as the difficulty of
maintaining large adult M. pyrifera in culture. This
section will specifically assess the culturing of early life
stages for experimental purposes.
Experimentation may begin from as early as the
sporulation stage when sorus tissue is exposed to differ-
ent experimental conditions, for these types of experi-
ments, sporulation can take place as advised above while
manipulating the factors of interest. After sporulation
takes place and before settlement occurs it is important
to achieve the desired concentration of spores for any
ongoing experiment, this will vary depending on the
needs of the experiment. Although a wide range of
spore densities have been applied in other studies
(Barrento et al., 2016; Buschmann et al., 2006, 2004;
Camus & Buschmann, 2017; Cie & Edwards, 2008;
Deysher & Dean, 1984; Leal et al., 2014; Macchiavello
et al., 2010; Muñoz et al., 2004; Westermeier et al.,
2006), the optimal range of settled spores for maximum
sporophyte production is roughly 50 spore mm
–2
(Reed,
Neushul, & Ebeling, 1991).
Once the desired concentration of spores is achieved,
the spore solution can be seeded to suitable culture
vessels depending on experimental purposes. Most stu-
dies utilize glass or plastic vessels, usually vials, flasks or
Petri dishes (Barrento et al., 2016; Carney & Edwards,
2010; Cie & Edwards, 2008; Deysher & Dean, 1984;
Hollarsmith et al., 2020; Morris et al., 2016;
Westermeier et al., 2006). When selecting vessels,
ensure they are of appropriate size, consider that settle-
ment is greatest on horizontal as opposed to vertical
surfaces (Reed et al., 1992) and if the experiment
requires observation of development, the vessel has
a flat, optically clear and uniform base. Observation is
best achieved using an inverted microscope with
a magnification range of 4x–80x depending on develop-
mental stage (Carney & Edwards, 2010; Hollarsmith
et al., 2020; Leal et al., 2017; Morris et al., 2016;
Muñoz et al., 2004).
Spore settlement generally ranges from 30 min to 1 h
in high light i.e., <50 µmol m
–2
s
–1
or 1–2 hrs in dark-
ness if the medium is not constantly agitated (Amsler &
Neushul, 1989, 1990). This will allow for the settlement
of the majority of viable spores. After 12–24 hrs, the
medium can be gently decanted and replaced to discard
any unsettled spores. Experimental treatments can com-
mence from this point if not already initiated from the
sporulation stage. If it is desirable that development
occurs in a free-floating state, i.e., not attached to the
bottom of the vessel, then aeration can be applied
through an airstone or through mechanical mixing
using a shaker table or stirring device. Depending on
experimental conditions, the development from spore
to juvenile sporophyte may progress over 2 to 4 weeks
(Camus & Buschmann, 2017; Carney & Edwards, 2010;
Hollarsmith et al., 2020; Neushul, 1963). If conditions
are not suitable for development, then development
may cease at a particular life stage and remain in
a vegetative state or perish (Lüning & Neushul, 1978;
Westermeier et al., 2006).
When performing experiments that seek to quantify
population characteristics rather than those at the indi-
vidual level, it is important to utilize sori from many
individual sporophytes (>30) as the response of
M. pyrifera at an individual level can be highly variable
due to plastic responses, genetic diversity and/or local
adaptations (i.e., Fernández, Navarro, Camus, Torres, &
Buschmann, 2021). It should be noted that each life
stage will respond differently to experimental condi-
tions, and this should be considered when designing
experiments.
Breeding
The purpose of breeding programs is generally to
enhance the performance of the macroalgal culture
(e.g., to increase yield, breed stress tolerance and
increase reproduction). This can be achieved by 1)
cross-breeding males and females that exhibit desirable
traits to create a hybrid generation, these are usually
identified from physiological experiments, 2) creating
a combination between hybridization and simple phe-
notypic mass selection or 3) more advanced gene editing
of particular individuals to promote certain character-
istics (Goecke, Klemetsdal, & Ergon, 2020).
For M. pyriferabreeding strategies are relatively
underdeveloped with most approaches focusing on
hybridization across one generation (Camus et al.,
2021; Murúa, Patiño, Müller, & Westermeier, 2021;
Raimondi, Reed, Gaylord, & Washburn, 2004;
Westermeier, Patiño, Müller, & Müller, 2010, 2011).
However, for other species that have a longer history
374 D. M. LE ET AL.

in commercial aquaculture, such as Saccharina spp.
and Undaria spp.,Pyropia spp., more advanced breed-
ing programs have been developed and have observed
success in increasing productivity, disease resistance,
heat tolerance, seasonal duration, and chemical con-
tents (Goecke et al., 2020; Hu et al., 2021; Hwang,
Yotsukura, Pang, Su, & Shan, 2019; Wang, Yao,
Zhang, & Duan, 2020). It is likely that as M. pyrifera
is incorporated into greater commercial production,
similar breeding programs will be undertaken, but this
is currently an area for further research. The selection
of particular traits will also benefit restoration efforts,
which is discussed in more detail below.
Farming
Macroalgal aquaculture is rapidly growing, driven by
the demands of the food, pharmaceutical, agricultural
and aquacultural industries (Ferdouse, Holdt, Smith,
Murúa, & Yang, 2018; Wernberg et al., 2019; Naylor
et al., 2021). M. pyrifera, being one of the fastest growing
macroalgal species with high biomass production (Reed
et al., 2008), high affinity for nutrients (Correa et al.,
2016; Purcell-Meyerink et al., 2021) and polysaccharide
content (Ortiz et al., 2009), holds significant value as
a desirable culture species.
The history of M. pyrifera farming provides many
examples of the challenges faced by this industry. Over
time, reliable protocols for mass production have been
developed and are now being implemented
(Buschmann et al., 2004; Westermeier et al., 2006;
Gutierrez et al., 2006; Macchiavello et al., 2010; Camus
et al., 2018b). Typically, culture and production rely on
sourcing spores directly either from wild populations or
from gametophytes, which are held under controlled
culture conditions. The latter approach allows the selec-
tion of specific traits (i.e., high-yield sporophytes)
through cross-breeding specific gametophytes
(Westermeier et al., 2010, 2011; Camus, Faugeron, &
Buschmann, 2018a; Buschmann et al., 2020; Camus
et al., 2021; Murúa et al., 2021). The set-up and require-
ment for growing kelp is discussed in two separate
phases: hatchery and grow-out.
Hatchery requirements
The purpose of this section is to provide a general over-
view of the essential capabilities a hatchery must sup-
port. Several very detailed guides on the requirements
for macroalgal hatcheries have been written (Andersen,
2005; Redmond, Green, Yarish, Kim, & Neefus, 2014).
Although we focus on M. pyrifera, the requirements will
scale well to many other laminarian kelps. Depending
on the scale of aquacultural application, hatchery
requirements will vary; however, approaches to produ-
cing and maintaining stock will be comparable regard-
less of scale. As with all culturing practices, sterility is
essential. All water should be filtered and sterilized to
the requirements laid out above, and air sources should
also be filtered using a 0.2 µm (polytetrafluoroethylene)
inline air filter.
Seeding stock onto seed lines is the most common
and by far the most efficient means for large-scale aqua-
culture production (Camus et al., 2018b). These seeded
lines should be maintained in the hatchery until juvenile
sporophytes of desirable size are present before being
relocated to an aquaculture setup in a natural environ-
ment where the grow-out phase will take place.
Preferable seed lines are synthetic twines such as nylon
and vinylon ~1–2 mm thick (Camus & Buschmann,
2017; Gutierrez et al., 2006); this should be wrapped
tightly around a sterilized PVC tube cut to an appro-
priate length for the application needed and the length
should match the depth of the aquaria that will house
the seed line as growth progresses (Camus &
Buschmann, 2017; Gutierrez et al., 2006). Seed lines
can be pre-prepared, wrapped in plastic wrap and stored
in a refrigerator for later use (Redmond et al., 2014).
Seed stock will either be from wild sourced repro-
ductive material or from gametophyte cultures already
held in the hatchery under vegetative conditions. If wild
sourced stock is used, then sporulation should take place
as described above whereby spores are released into
a glass vessel after the pretreatment phase, and
a density of ~40 000 spores ml
–1
is desirable (Camus &
Buschmann, 2017; Gutierrez et al., 2006). Spore solution
is added to aquarium or culture containers, previously
filled with filtered seawater, holding the PVC tubes
wrapped with seed line (Camus & Buschmann, 2017;
Redmond et al., 2014). These should be left for 24 hrs
before removing the seed line wrapped tubes and pla-
cing them into culture aquaria tanks filled with
Provasoli enriched seawater held at a constant tempera-
ture. If gametophyte stock cultures are used, then these
should be vegetatively propagated in order to obtain
a high density of gametophytes. This is done under
similar culture conditions described in the germplasm
banking section above. This may take weeks to months
depending on the initial density of gametophytes held in
the culture (Barrento et al., 2016; Westermeier et al.,
2006). Once a dense culture is present, it can be trans-
ferred to an electric blender or a tissue disruptor and
fragmented until a dark brown solution results. This
solution can then be sprayed directly onto the seed
line using a pressure sprayer, left for 30 min – 1 hr,
before being placed in culture aquaria tanks. Seawater
changes should be carried out weekly to refresh nutrient
APPLIED PHYCOLOGY 375

concentrations (Redmond et al., 2014). Care should be
taken to avoid overstocking aquaria with seed line PVC
tubes as this can create shading, limit water movement
and deplete nutrients, meaning water will need to be
changed more frequently. Camus and Buschmann
(2017) detail the optimal production conditions for
rapid growth at a temperature of 12°C, a photon flux
density of 12 µmol m
–2
s
–1
, a photo period of 16 L:8D
and aeration of 414 l h
–1
. Under these conditions they
were able to produce 4–5 mm long juvenile sporophytes
after 45 days post sporulation. During the hatchery
phase, environmental conditions should be monitored
daily to ensure optimal conditions for growth.
Another applicable seeding approach is to attach
free-floating sporophytes directly onto seed lines.
This method, which has been described in detail
elsewhere (see Camus, Infante, & Buschmann,
2018b; Westermeier et al., 2006), is less common
than direct seeding due to its labour-intensive nature
(Camus et al., 2018b). Briefly, embryonic sporophytes
are produced in floating cultures, by providing con-
stant water motion in the culture vessel, until
a desired size of 4–10 cm is reached. These are
then extracted from the culture and sown directly
onto seed lines either using adhesives or simply by
inserting the holdfast between the weaves of the seed
lines. Individuals are typically spaced 10–30 cm apart
for a grow-out.
Grow-out requirements
The grow-out phase should typically be carried out in
a natural environment where nutrient and light availabil-
ity are not limiting, where water motion is present but not
detrimental and where temperature ranges are well within
the thermal limits of M. pyrifera (Fain & Murray, 1982;
Deysher & Dean, 1984; Harrison & Hurd, 2001; Camus &
Buschmann, 2017; Camus et al., 2018; North et al., 1986).
A wide range of approaches have been trialled using
varying infrastructures, ranging from shallow-water grid
arrays to offshore deep-water suspension lines
(Buschmann et al., 2014; Bak et al., 2018; Bak,
Gregersen, & Infante, 2020; Camus et al., 2018b). Grow-
out facilities are usually located in shallow coastal envir-
onments, <30 m deep with strong current flow, nitrogen
levels of no less than 5–8 µMol and high light availability.
Site selection is a critical decision as high-temperature
pulses (normally not detected by observing average values,
salinities and nutrient drops and pest organism (epibionts,
grazer and eventually pathogens) can determine the via-
bility of the farming activity (Camus et al., 2018b).
Grid arrays are commonly employed as grow-out
structures (Bak et al., 2020) as they provide the great-
est area coverage and are rigid and maintain a stable
desired depth (Fig 3). These arrays involve a series of
mooring blocks or steel anchors that fix the grid to
the seabed. From these moorings, ropes are sus-
pended in a cross-linked fashion creating
a semirigid grid (Camus et al., 2018b). In each grid,
backbone lines can be strung from one side to the
other, parallel to one another, and it is on these
backbone lines that the seed line will be deployed.
Seed line (either direct seeded or free-floating) can be
deployed by feeding one end of the backbone line
through the PVC seed line tube, tying the end of
the seed line to the backbone line, attaching it to
the grid and then moving the PVC seed line tube
Figure 3. Culture infrastructure for the production of Macrocystis pyrifera. Seed line in hatchery aquaria (a), layout of grow-out grid
(modified from Camus et al., 2018b) (b), seed line attachment to main backbone line (modified from Gutierrez et al., 2006) (c), low
angle view of grid array (modified from Bak et al., 2020) (d).
376 D. M. LE ET AL.

along the backbone line, letting it coil around the
backbone as it moves along, before tying it off once
it reaches the other end (Figure 2).
Setting the depth of the grid array is important
(Camus et al., 2018b) and should be based on knowledge
of light, nutrient, temperature, and water motion char-
acteristics at the chosen site. The depth of the array can
be manipulated by attaching surface or subsurface
buoys to the grid to provide floatation while altering
the length of the mooring ropes that suspend the grid.
Manipulation using buoys initially attached to the grid
is essential to maintain buoyancy and as pneumatocysts
develop, buoys can be removed and weights added to
offset the floatation of the developing kelp.
Seasonality dictates growth and for M. pyrifera grow-
out will typically be most successful if commenced in
autumn/winter with a spring/summer harvest
(Buschmann et al., 2004, 2014; Camus et al., 2018b). This
will, however, depend on local conditions and in situ phy-
siological understanding (Gerard, 1982; Varela et al.,
2018).
Restoration
Significant declines in kelp forest habitat have been
recorded across the globe over the past 50–
100 years (Desmond et al., 2015; Friedlander et al.,
2018; Krumhansl et al., 2016; Layton et al., 2020;
Wernberg et al., 2019), included in that loss have
been large swaths of M. pyrifera (Hay, 1990;
Johnson et al., 2011; Butler, Lucieer, Wotherspoon,
& Johnson, 2020; Layton et al., 2020, Tail et al.,
2021). The driving factors behind kelp forest decline
are relatively well understood and include increas-
ing sea surface temperature (Filbee-Dexter et al.,
2016; Johnson et al., 2011; Wernberg et al., 2013),
light limitation (Cie & Edwards, 2008; Desmond
et al., 2015; Deysher & Dean, 1984; Fain &
Murray, 1982; Navarro, Mansilla, & Palacios,
2007), increased storm frequency (Buschmann
et al., 2004), sedimentation (Connell, 2003; Filbee-
Dexter & Wernberg, 2018), food web alterations
through overfishing (Friedlander et al., 2018;
Krumhansl et al., 2016), eutrophication (Filbee-
Dexter & Wernberg, 2018) and direct harvest
(Buschmann et al., 2014). This decade, 2021–2030,
is the UN Decade on Ecosystem Restoration with
a goal to restore 350 million hectares of degraded
ecosystems (FAO, 2020b). Arguably, the revitaliza-
tion and restoration of vast areas of lost kelp forest
would deliver some of the greatest benefits in terms
of fisheries productivity, ecosystem services and
socio-ecological outcomes of any ecosystem type.
Early kelp forest restoration efforts began sometime
before 1970 and have shown mixed results over the last
five decades (Campbell, Marzinelli, Coleman, Verge, &
Steinberg, 2014; Carney, Waaland, Klinger, & Ewing,
2005; Fredriksen et al., 2020; Mcleod et al., 2018;
Mearns, Hanan, & Harris, 1997; North, 1976;
Westermeier et al., 2014; Wood et al., 2019).
Approaches have included encouraging natural recruit-
ment through the placement of artificial substrate, to
directly seeding habitat with microscopic life stages, to
transplantation of juvenile and adult individuals. Many
of these approaches have been extremely time, labour
and financially demanding, often making them prohibi-
tively expensive to maintain. As approaches advance,
a move to aquacultural practices (Alleway et al., 2019;
Froehlich, Gentry, & Halpern, 2017; Giangrande,
Gravina, Rossi, Longo, & Pierri, 2021) is emerging
which allows large quantities of stock to be produced
under laboratory conditions with known genetic and
physiological characteristics. This stock can then be out-
planted into the receiving environment when condi-
tions are appropriate. Out-planting approaches vary,
and their success is highly dependent on the site condi-
tions. This area of research is currently underdeveloped
but promising results have been seen using seed line
attached directly to or above the substrate (Giangrande
et al., 2021; Kraufvelin & Díaz, 2015), seeded tiles
attached to the seafloor (Layton et al., 2020; Shelamoff
et al., 2020) and an approach termed “green gravel”
which is gravel seeded in the laboratory and distributed
directly onto the seafloor (Coleman et al., 2020;
Fredriksen et al., 2020). All of these approaches out-
plant when M. pyrifera are at the juvenile sporophyte
stage.
Conclusion
This critical analysis summarizes the current state of
knowledge regarding the culture of M. pyrifera and
details the necessary steps for obtaining and preserving
stock as well as propagation for experimental, aquacul-
tural and restorative purposes. Current threats to
M. pyrifera and the value this species holds from an
aquacultural perspective mean that a concerted effort
is required to standardize preservation and propagation
methods. It is essential to safeguard this iconic species
from the current local and global stressors and to lock-
APPLIED PHYCOLOGY 377

in the many benefits it offers for sustainable
aquaculture.
Acknowledgments
This study was funded by New Zealand Ministry of Business,
Innovation and Employment to MJD and CDH (grant number
UOOX1908 - “Cultivating resilient marine forests to rebuild
productive coastal ecosystems”). DML was funded by
a postgraduate scholarship from University of Otago, New
Zealand.
Authors’ contribution
CDH and MJD proposed the idea behind this study. DML and
MJD wrote most parts of the manuscript as well as produced
all the figures. However, all authors contributed critically to
the draft with valuable inputs and experience. All authors read
and gave final approval for publication.
Disclosure statement
No potential conflict of interest was reported by the author(s).
Funding
This work was supported by the Ministry of Business,
Innovation and Employment, New Zealand [UOOX1908].
ORCID
Duong M. Le http://orcid.org/0000-0001-8424-4624
Alejandro H. Buschmann http://orcid.org/0000-0003-
3246-681X
Catriona L. Hurd http://orcid.org/0000-0001-9965-4917
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