doi: 10.1007/978-3-030-23459-1

Chapter 8

The Cricket Gryllus bimaculatus:

Techniques for Quantitative and Functional

Genetic Analyses of Cricket Biology

Arpita Kulkarni and Cassandra G. Extavour

Abstract All extant species are an outcome of nature’s “experiments” during

evolution, and hence multiple species need to be studied and compared to gain a

thorough understanding of evolutionary processes. The ﬁeld of evolutionary devel-

opmental biology (evo-devo) aspires to expand the number of species studi ed,

because most functional genetic studies in animals have been limited to a small

number of “traditional” model organisms, many of which belong to the same phylum

(Chordata). The phylum Arthropoda, and particularly its component class Insecta,

possesses many important characteristics that are considered favorable and attractive

for evo-devo research, including an astonishing diversity of extant species and a

wide disparity in body plans. The development of the most thoroughly inves tigated

insect genetic model system to date, the fruit ﬂy Drosophila melanogaster

(a holometabolous insect), appears highly derived with respect to other insects and

indeed with respect to most arthropods. In comparison, crickets (a basally branching

hemimetabolous insect line age compared to the Holometabola) are thought to

embody many developmental features that make them more representative of

insects. Here we focus on crickets as emerging model s to study problems in a

wide range of biological areas and summarize the currently available molecular,

genomic, forward and reverse genetic, imaging and computational tool kit that has

been establis hed or adapted for cricket research. With an emphasis on the cricket

species Gryllus bimaculatus, we highlight recent efforts made by the scientiﬁc

community in establishing this species as a laboratory model for cellular biology

and developmental genetics. This broad toolkit has the potential to acceler ate many

traditional areas of cricket research, including studies of adaptation, evolution,

A. Kulkarni

Department of Organismic and Evolutionary Biology, Harvard University, Cambridge, MA,

USA

C. G. Extavour (

Department of Organismic and Evolutionary Biology, Harvard University, Cambridge, MA,

USA

Department of Molecular and Cellular Biology, Harvard University, Cambridge, MA, USA

e-mail: [email protected]

W. Tworzydlo, S. M. Bilinski (eds.), Evo-Devo: Non-model Species in Cell and

Developmental Biology, Results and Problems in Cell Differentiation 68,

https://doi.org/10.1007/978-3-030-23459-1_8

183

neuroethology, physiology, endocrinology, regeneration, and reproductive behavior.

It may also help to establish newer areas, for example, the use of crickets as animal

infection model systems and human food sources.

8.1 Introduction

All cellular life forms share a last common ancestor. Every extant species is a current

outcome of an evolutionary process that has taken place over hundreds of millions of

years. Thus, each species that exists today can be used as a data point toward

increasing our understanding of the living world. Comparative study of the devel-

opmental biology of multicellular organisms motivates the ﬁeld of evolutionary

developmental biology, or “evo-devo.” However, to date only a relatively small

number of species has been used to study the functions of genes that regulate

animal developmental processes, which limits our understanding of how develop-

mental genetic changes underpin evolution. If the idealized goal of evo-devo

research is to do a speci es comparison that includes representatives of all major

evolutionary transitions, then this calls for the establishment of many more animals

as laboratory model organisms than is currently the case. As a step in this direction,

developmental biologists are increasingly choosing new animal models that are

suitable to address thus far neglected areas of research, while sim ultaneously

selecting these candidates for their ability to fulﬁll other criteria relevant for

evo-devo work. Some criteria that could maximize the scientiﬁc gains achieved by

establishing new models include choosing organisms that belong to clades that are

representative of a wide range of ecolog ical niches, and those belonging to phyla that

are species-rich, display high diversity in form and function, have interdisciplinary

scientiﬁc appeal, are economical to maintain in the laboratory, and could inform

issues that directly affect humans (e.g., disease or agriculture). Strategically choos-

ing and studying examp les satisfying some or all of these criteria may therefore be

impactful, evolutionarily informative, and a good use of limited resour ces.

The phylum Arthropoda contains multiple species that satisfy many of the above

criteria and has thus played a prominent role in modern evo-devo research. Impor-

tantly, good fossil records exist for this phylum, providing researchers with snap-

shots into the evolutionary past and aiding in comparative work. For example, the

EDNA fossil insect database lists over 23,000 species (Giribet and Edgecombe

2013; Mitchell 2013), with the earliest records of arthropod fossils dating back to

nearly 555 million years ago (Mya) during the Cambrian era (Harvey et al. 2012;

Zhang et al. 2010; Vaccari et al. 2004). Undoubtedly, access to such fossil records is

essential to understanding the key phenotypic innovations that have made

Arthropoda species-rich and evolutionarily successful (Mayhew 2007).

Extensive studies on insects, which account for the majority of all species

described on earth (Wheeler 1990; Grimaldi and Engel 2005) and for ~85% of all

arthropod diversity (Giribet and Edgecombe 2013), have pioneered and shaped the

evo-devo ﬁeld. This work has informed us of the genetic and evolutionary basis of

184 A. Kulkarni and C. G. Extavour

pivotal developmental mechanisms. For example, the description of the ﬁrst home-

otic mutant (Bridges and Morgan 1923), and the realization of the signiﬁcance of

conserved developmental genes in body patterning and in the evolution of different

body plans across animals, come from studies in the insect Drosophila melanogaster

(Lewis 1978; Nüsslein-Volhard and Wieschaus 1980; Duboule and Dolle 1989;

Graham et al. 1989 ; Panganiban et al. 1997; Heffer et al. 2013). Other insect-based

research that has broadened our understanding of evolutionary processes includes

work on key evolutionary innovations such as the insect body plan (Grimaldi and

Engel 2005), metamorphosis (Truman and Riddiford 1999), development of wings

(Nicholson et al. 2014; Alexander 2018; Bruce and Patel 2018 ; Linz and Tomoyasu

2018), morphological novelties (Kijimoto et al. 2013), and insect eusociality (Toth

and Rehan 2017). The widespread scope of such research is a result of the practical

advantages that come with working on insects: insect phylogeny is well established

(Giribet and Edgecombe 2013), many species are easy to mai ntain and culture in the

laboratory, are often amenable to functional genetic analysis, in many instances

produce easily accessible large broods suitable for external manipulation, and often

have life cycles sufﬁciently short to allow laboratory rearing and multigenerational

analysis.

To date, popular insect models to study the genetic basis of development have

included the fruit ﬂy Drosophila melanogaster (Diptera) (Demerec 1950), the ﬂour

beetle Tribolium castaneum (Coleoptera) (Sokoloff 1966, 1972, 1974, 1977; Denell

2008), the honeybee Apis mellifera (Hymenoptera) (Gould and Grould 1995;

Oldroyd and Thompson 2006), the wasp Nasonia vitripennis (Hymenoptera)

(Werren and Loehlin 2009), and the silk moth Bombyx mori (Lepidoptera) (Xia

et al. 2004; Goldsmith et al. 2005; Meng et al. 2017). All of these insects, however,

share a commonality: they belong to the same insect superorder of Holometabola, or

insects that undergo complete metamorphosis during development. Complete meta-

morphosis is characterized by a pupal stage in the transition from larvae to adults,

with neither the larval nor the pupal stages resembling the ﬁnal adult form (Truman

and Riddiford

1999). Such insects display a number of evolutionarily derived

developmental characters that are not generally representative of all insects, let alone

all arthropods (Mito and Noji 2008). To correct the overrepresentation of holome-

tabolous insects in modern comparative developmental literature, insects branching

basally to the Holometabola should be studied, as they, based on parsimony, appear

to display characters that are likely ancestral to insects and in some cases also to

arthropods. These insects are the Hemimetabola, insects displaying incomplete

metamorphosis and lacking pupal stages during development. The embryo in such

insects develops into a miniature adult (referred to as a nymph or a juvenile) which

then undergoes several successive molts before reaching a dulthood and sexual

maturity. Orthopterans (crickets, grasshoppers, and locusts), one of the most abun-

dant and dominant terrestrial insect groups, are in this category (Grimaldi and Engel

2005). Orthopterans display extraordinary diversity in developmental processes and

are also economical ly important herbivores, which has resulted in them becoming

popular for functional genetic research in recent years.

8 The Cricket Gryllus bimaculatus: Techniques for Quantitative .. . 185

For the rest of this chapter, we focus on crickets as promising evo-devo models,

deserving of serious regard. We discuss Gryllus bimaculatus,aﬁeld cricket species,

introduce this model system, and discuss recent advances in estab lishing this animal

for cell, developmental, and genetic research.

8.2 A Hemimetabolous Insect Model: The Cricket Gryllus

bimaculatus De Geer

G. bimaculatus is a cosm opolitan orthopteran belonging to the family Gryllidae and

is, to our knowledge, the most widespread of all Gryllus specie s for laboratory

studies (Otte and Cade 1984). Although its use for developmental genetics is

relatively recent, this species is by no means new to biological research:

G. bimaculatus has been extensively used to inform areas such as neurobiology ,

insect physiology, reproduction, and behavior since the 1960s (Huber et al. 1989;

Engel and Hoy 1999; Paydar et al. 1999; Wenzel and Hedwig 1999; Hedwig and

Poulet 2004; Nakamura et al. 2008a, b; Horch et al. 2017b). The discovery of RNA

interference (RNAi) as a mechanism for abrogating gene function (reviewed by Sen

and Blau 2006) has greatly accelerated G. bimaculatus research (Mito et al. 2011),

yielding important information about the developmental biology of this organism

and unveiling its potential as an upcoming functional genetics laboratory model.

This specie s was ﬁrst described by Baron Charles de Geer in 1773 (Geer 1773)

and named Gryllus (meaning “ cricket” in Latin) bimaculatus (from the Latin “mac-

ula” for “spot ” ). Indeed, this species is commonly referred to as the “two spotted

ﬁeld cricket,” for the white spot that this species displays on the dorsal surfa ce of the

forewings next to the pronotal margin (Fig. 8.1a) (Otte and Cade 1984). Some of the

morphological keys used to distinguish adult G. bimaculatus from other similar

looking ﬁeld cricket speci es, both within and outside the genus Gryllus (Otte and

Cade 1984), include the white spots, a black colored adult body size of ~3 0 mm

lacking any bands, forewings nearly covering and large hindwings extending well

beyond the abdomen, and an ovipositor (in females) slightly longer than the hind

femora (Fig. 8.1a). As an additional tool in the ﬁeld, entomologists have documented

that G. bimaculatus does not undergo an obligator y or facultative winter diapause

(i.e., developmental arrest in response to adverse environmental conditions such as

temperature and/or photoperiod), at any stage of its life cycle (Bigelow 1962). Such a

lifestyle is called a homodynamic life cycle, and is in contrast to the heterodynamic

lifestyle observed in many other Gryllus species entering diapause. Examples of

overwintering species in this genus include G. pennsylvanicus

, G. campestris,

G. fultoni, G. veletis, G. vernalis, and G. ﬁrmus (Bigelow 1962). This means that

it is possible to collect G. bimaculatus in the ﬁeld across seasons and makes it easy to

culture and breed this species in the laboratory all year-round. Additionally, in the

ﬁeld, G. bimaculatus is often found close to human settlements, on the ground

surface or in soil cracks (Otte and Cade 1984).

186 A. Kulkarni and C. G. Extavour

While one could collect this species from the wild, the easiest way to establish a

laboratory culture of G. bimaculatus is from animal pet suppliers. Multiple online

pet suppliers based in various countries (e.g., Pets at Home, UK; Bugs International,

Germany) culture and distribute this species as live crickets for captivity feeding. It

is important to note, however, that at the time of writing, the United States is an

exception: the United States Department of Agriculture (USDA ) does not permit

commercial distribution of G. bimaculatus in the United States, and the cricket spe-

cies commonly commercially available for purchase are Gryllodes sigillatus and

Acheta domesticus (examples of online retailers selling cricket species in the United

States at the time of writing include Ghann’s Crickets, Top Hat Cricket Farm,

Fluker’s Cricket Farm, and Premium Crickets). Being bulkier and meatier than

other cricket species (Fig. 8.1b dorsal and lateral view), G. bimaculatus is

Fig. 8.1 G. bimaculatus morphology. (a) G. bimaculatus adult female (left), adult male (center),

and male nymph (right) displaying some morphological keys used to identify this species. The

characteristic white spots (marked with white asterisks on the adult female and male) on the

forewings next to the pronotal margin are shown. Animals are black in color and are therefore

vernacularly also known as the “black cricket” in some parts of the world. The body (in both sexes)

lacks any bands of contrasting pigmentation; forewings nearly cover the abdomen, and hindwings

extend well beyond the abdomen; females possess an ovipositor (marked with pink asterisk), used

to deposit eggs. The presence or absence of an ovipositor can be used to sex animals in nymphal

stages (developmental stages prior to becoming an adult). Note: The adult female and male

photographed in this picture have broken left antennae; normally both antennae are of similar

lengths. Scale bar is 1.5 cm. (b) Comparison (dorsal and lateral view) between an adult G. assimilis

(unmarked) and G. bimaculatus female (marked with white asterisk). Note the brown body color,

leaner and less bulky adult body, and the absence of the white spots on the forewings in G. assimilis.

images of the cricket head (front view) showing the white eye color in these mutants (left) compared

to wild-type pigmented eyes shown in (e) (right)

8 The Cricket Gryllus bimaculatus: Techniques for Quantitative .. . 187

preferentially exploited in multiple countries as an inexpensive food source for

humans (described below, Sect. 8.8) and for insectivorous animals housed in

captivity (Mito and Noji 2008). This has signiﬁcantly raised their potential economic

importance in recent years. Because G. bimaculatus is quite heat tolerant and one of

the very few insects that can be reared at 37



C (human body temperature), this

species is also promising as a simple animal infection model system and has found

application in studying human pathogenic bacteria (e.g., Staphylococcus aureus,

Pseudomonas aeruginosa, and Listeria monocytogenes) (Kochi et al. 2016) and

fungi including various Candida speci es (Kochi et al. 2017). Epizootic viral diseases

are devastating in crickets (and for cricket-rearing facilities), wiping out entire

colonies and becoming difﬁcult to eradicate. Researchers who wish to culture

G. bimaculatus, therefore, should be aware that this species is susceptible to the

G. bimaculatus nudivirus (GbNV), known to infect nymphs and adults (Wang and

Jehle 2009) and the cricket iridovirus (CrIV) (Kleespies et al. 1999), but is reportedly

resistant to the cricket paralysis virus (CrPV ) and the potent A. domesticus

densovirus (AdDNV) (Szelei et al. 2011).

As a hemimetabolous insect, G. bimaculatus displays a short germ band during

embryonic development and thus differs substantially from the well-studied long

germ band characteristic of Drosophila. Short germ band development refers to a

form of insect body patterning that is thought to be ancestral to arthropods (reviewed

by Davis and Patel 2002) and present in many extant insects including crickets. In

this form of development, only the anterior body segments (head only or the head

and thorax) are speciﬁed in the early embryonic rudiment before gastrulation,

whereas posterior segments (the thorax or the thorax and abdomen) are formed

sequentially later in development during a secondary growth phase (reviewed by

Krause 1939; Davis and Patel 2002; Liu and Kaufman 2005). In contrast, insects

such as Drosophila follow the presumed derived long germ band type of develop-

ment, whereby all segm ents are speciﬁed near simultaneously during the early

blastoderm stage (Krause 1939; Campos-Ortega and Hartenstein 1985; Lohs-

Schardin et al. 1979; Liu and Kaufman 2005). Another way in which crickets may

display putative ancestral insect characteristics is in the structure of its ovaries. The

G. bimaculatus ovary is panoistic, meani ng that there are no germ-line-derived nurse

cells that provide cytoplasmic content to growing oocytes (Büning 1994). Instead,

every germ-line cell (i.e., every cystoblast) in the adult female ovary is thought to

give rise to an oocyte (Büning 1994). In contrast, in the meriostic type of ovaries, as

seen in Drosophila and nearly all other holometabolous insects (for details see

Bilinski et al. 2017), the oocytes are connected to groups of germ-line cells called

nurse cells. Panoistic ovary type and short germ development are, based on parsi-

mony, thought to be features ancestral to insects and possibly to Pancrustacea.

Consequently, it has been proposed that this species has the potential to serve as a

representative study model for basally branching, hemimetabolous insect and arthro-

pod lineages (Sander 1997; Mito and Noji 2008).

Sex determination in G. bimaculatus is thought to follow the XX/X0 system, with

females being the homogametic sex and having a chromosome complement of

2n ¼ 28 + XX (Yoshimura 2005) and having a predicted genome size of a

188 A. Kulkarni and C. G. Extavour

few gigabases (Mito and Noji 2008). G. bimaculatus is polyandrous—females are

known to mate with several males and exert postcopulatory mate choice (Tregenza

and Wedell 1998). This polyandry is associated with increased egg-hatching rates

and is hypothesized to prevent effects of inbreeding in wild populations (Simmons

1986, 1987; Tregenza and Wedell 1998, 2002; Bretman and Tregenza 2005).

Indeed, G. bimaculatus females can lay many hundreds or thousands of eggs over

their lifetime in the lab and, in our hands, have been maintained successfully as an

inbred line (originally founded from a few dozen individuals) for over a decade,

without any noticeable decline in health that might be attributed to inbreeding

depression (Extavour lab, unpublished observations).

8.3 Cricket Sources, Animal Husbandry, Life Cycle,

and Available Strains

At the time of writing and to the best of our knowledge (as described above), most

current working laboratory cultures of G. bimaculatus were either established from

adults purchased from commercial vendors (e.g., Tsukiyono Farm, Gunma, Japan;

Scope Reptile Pet Store, Okayama, Japan; Livefood UK Ltd., UK; Kreca Ento-Feed

BV, the Netherlands) or caught in the wild. However, this genus contains many

species that are morphologically very similar, many species are know n to overlap

with G. bimaculatus in their local distribution, and adequate species-level, promi-

nent morphological keys are lacking within this genus. Although some keys have

been described (e.g., Nickle and Walker 1974), including the morphological char-

acters described above, to an inexperienced eye, many of these features are often

distinguishable only in comparison with another species present, or are easier to

observe in preserved specimens than in live animals. We thus recommend

performing molecular barcoding (e.g., using 16s ribosomal DNA or the cytochrome

b mitochondrial DNA sequence) of the founding adults of a new colony, whether

purchased commerci ally or captured in the wild, to ensure that all founding adults

are indeed G. bimaculatus (Ferreira and Ferguson 2010).

Rearing G. bimaculatus is straightforward, and detailed cricket husbandry pro-

tocols are well described for this species (Mito and Noj i 2008; Kainz et al. 2011;

Kochi et al. 2016). Crickets (nymphs and adults) can be kept as inbred lines at

26–30



C in well-ventilated plastic cages with egg cartons (Fig. 8.2a) or crumpled

paper for shelter, and can be maintained on either a 12 h light/12 h dark (Kainz et al.

2011) or a 10 h day/14 h dark photoperiod (Mito and Noji 2008). They can be fed on

general insect food or artiﬁcial insect diets (e.g., Oriental Yeast Co., Ltd., Tokyo,

Japan), artiﬁcial ﬁsh food, ﬁnely ground dry cat food (e.g., Purina Kitten Chow), a

mixture of oils and whole grain cereals (Kainz et al. 2011), or a combination of these

food sources (Fig. 8.2b). Cricket Quencher water gel (Fluker Farms) can be used as a

water source. Alternatively, a 50 mL falcon tube ﬁlled with water and stopped with

cotton, or wet tissue or cotton in petri dishes, can also serve as water sources

8 The Cricket Gryllus bimaculatus: Techniques for Quantitative .. . 189

(Fig. 8.2 b ). Crickets will oviposit fertilized eggs into damp sand (e.g., Sandtastik

Sparkling White Play Sand, Product Code PLA0050), wet paper towels, Whatman

paper, or wet cotton placed in petri plates in cricket cages (Fig. 8.2b inset, d). These

eggs will develop successfully and hatch in 12–14 days (Fig. 8.2c) under the

following conditions: incubation at ~28–29



C with 70% humidity, dead or moldy

Fig. 8.2 G. bimaculatus husbandry. (a) (side view) and (b) (top view) showing a well-ventilated

plastic container used for housing a cricket colony. Note the use of egg cartons for providing shelter,

ground cat food, and a 50 ml Falcon ﬁlled with water and stopped with cotton as a food and water

source, respectively. A wet cotton plate (seen in a, b, and b inset) is placed in the adult cages for

females to oviposit fertilized eggs. Oviposited eggs (higher magniﬁcation shown in (d)) need to be

kept moist and clean until the eggs hatch. (c) A close-up of a cotton plate showing newly emerged

cricket hatchlings, which can then be transferred into new plastic cages with food, water, and shelter

until they reach adulthood. (e) Two G. bimaculatus eggs (6 days after egg laying) imaged under

bright ﬁeld white light (top) and green ﬂorescent light (bottom). Both eggs are progeny obtained

from a cross between the histone2B-GFP (H2B-GFP) transgenic and wild-type G. bimaculatus line.

Embryos carrying the H2B-GFP transgene (bottom egg marked with white asterisk) can be

distinguished from non-transgenic embryos (top egg) based on the presence of bright ﬂorescent

nuclei, which is detectable from day 5 until day 10 after egg laying

190 A. Kulkarni and C. G. Extavour

embryo removal on a regular basis, and maintenance of a moist and clean substrate

(Mito and Noji 2008; Donoughe and Extavour 2016). Embryonic development for

this species has been divided into 16 stages based on morphological features of

eggs, developing embryos and their appendages (Niwa et al. 1997; Donoughe and

Extavour 2016). After hatching, nymphs undergo eight nymphal molts to ﬁnally

become adults over the next 5 weeks. The generation time (total time to adulthood

and sexual maturity) of G. bimaculatus is thus approximately 7 weeks at 29



(Fig. 8.3). Adults are thought to reach maximum fecundi ty 1 week after the ﬁnal

molt (Mito and Noji 2008). Sexing and identiﬁcation of virgin males and females are

straightforward: late-stage male and female nymphs can be separated based on the

presence or absence of an ovipositor and then isolated until they undergo the ﬁnal

molt to sexual maturity (Fig. 8.1a). This also helps in setting up single mating

crosses, for example, to establish genetically modiﬁed lines. Similarly, precisely

timed egg collections are possible by placing egg collection petri dishes in the cages

and removing them at desired intervals (described in Donoughe and Extavour 2016).

Fig. 8.3 A schematic showing an overview of the G. bimaculatus life cycle. Selected embryonic

and nymphal developmental stages are shown, alongside the duration of each developmental stage

depicted in hours (orange arc and lines), days (green arc and lines), or weeks (blue arc and lines)

after egg laying. The colored arcs indicate the entire duration of time occupied by the indicated

developmental stage, whereas the lines show a cartoon schematic of the selected stages within this

time window. Each displayed embryonic stage during embryogenesis shows the position of the

embryo (in gray) relative to the yolk (yellow) within the egg and has a small description of features

that are characteristic of that developmental stage (depicted as egg stage “EgS”). Dotted lines with

arrowheads indicate the different movements that the embryo makes during the course of develop-

ment in this species. Upon hatching, nymphs undergo eight nymphal molts (indicated by solid black

arrows) to reach adulthood. Newly emerged adult animals are sexually mature at molting and begin

mating soon after. This ﬁgure is modiﬁed from Donoughe and Extavour (2016)

8 The Cricket Gryllus bimaculatus: Techniques for Quantitative .. . 191

A G. bimaculatus spontaneous mutant strain with white eyes (Fig. 8.1c–e) was

isolated by Isao Nakatani and colleagues at the University of Yamagata, Japan, in

1989 (referenced in Mito and Noji 2008). This is, to our knowledge, currently the

only available mutant, and its phenotype is caused by an autosomal recessive

mutation, referred to as gwhite (Niwa et al. 1997; Mito and Noji 2008). This mutant

is sometimes preferred for whole-mount gene expression analysis at late stages of

embryogenesis, owing to the fact that at this stage, the tissues are more transparent

compared to wild type (Niwa et al. 1997; Mito and Noji 2008).

8.4 Techniques for Quantitative and Functional Genetic

Analyses in G. bimaculatus

Here we discuss protocols and methodologies that have been established and are

currently available in the cricket G. bimaculatus, with the aim of making new users

aware of the plethora of techniques at their disposal. Detailed descriptions of these

published techniques and step-by-step protocols are thus avoided in this chapter

(we refer the reader to Horch et al. 2017a, b for detailed protocols). While these

protocols are now well established in crickets, these tools are not limited to them and

could in principle be modiﬁed or adapted for use in other hemimetabolous insects to

further species-speciﬁc research.

8.4.1 Precise Embryonic Staging System

To make meaningful observations of deviations from normal embryonic develop-

ment, one ﬁrst needs a wild-type reference for any given species. Donoughe and

Extavour (2016) have reported a detailed embryonic staging system for

G. bimaculatus (Figs . 8.4 and 8.5). This system is based on externally observable

characters of the developing cricket embryo that are visible through the eggshell,

thereby circumventing the need for embryonic disse ctions to ascertain embryonic

developmental stage. This is especially informative for studying early embryos of

insects such as crickets, which are embedded within a large amount of opaque yolk,

making direct observations through the eggshell difﬁcult if not impossible.

G. bimaculatus development, based on this staging, is presented as 24 “egg stages”

and encompasses the entire development of the animal from fertilization to hatching,

based solely on external observable egg characters (called stage identiﬁers). Each of

the 24 “egg stages” described here corresponds to one or more of 16 “ embryonic

stages.” For each stage, the authors provide a list of embryonic developmental

features deﬁning that stage, including features of body segmentation, mesoderm,

and appendage formation. Determining the stage of embryogenesis through the

eggshell is a useful complement to earlier described staging schemes that require

dissection of the embryo (Mito and Noji 2008; Kainz 2009).

192 A. Kulkarni and C. G. Extavour

schematicbright field

egg

stage

(EgS)

embryonic

stage (ES)

lateral view of egg or

ventral view of embryo

1.0 – 1.4

1.6 – 4.0

4.0 – 5.2

5.2 – 6.5

7.5 – 8.5

7.0 – 7.5

6.0 – 7.0

1.5

7.2 – 8.0

8.7 – 9.5

8.0 – 8.7

8.5 – 9.0

Fig. 8.4 A detailed description of the egg stages (EgS 1–12) in G. bimaculatus. Micrographs in

second column from left help display the morphological features of the egg that can be used to

assign embryos to an egg stage (EgS) and also describe the corresponding morphological features

and embryonic stage (ES) of the embryo within the egg. In the schematic, the embryo is depicted in

gray and yolk in yellow. Micrographs in the right second column from the right are not to scale, and

are taken using lateral views of either a H2B-GFP transgenic live embryo (EgS 1–5) or ventral

views of dissected and ﬁxed, Hoechst 33342-stained embryos (EgS 6-182). Micrographs are not to

scale. This ﬁgure is modiﬁed from Donoughe and Extavour (2016)

8 The Cricket Gryllus bimaculatus: Techniques for Quantitative .. . 193

schematicbright field

egg

stage

(ES)

embryonic

stage (ES)

lateral view of egg or

ventral view of embryo

9.0 – 9.9

11 – 12

13 – 14

12 – 13

11 – 12

12 – 14

Fig. 8.5 A detailed description of the egg stages (EgS 12–24) in G. bimaculatus. Staging system

for egg stages (EgS) 12–24 continued from Fig. 8.4. This staging system ends at hatching and does

not include postembryonic development of nymphs to adulthood. This ﬁgure is modiﬁed from

Donoughe and Extavour (2016)

194 A. Kulkarni and C. G. Extavour

8.4.2 Injection Methods for Eggs, Nymphs, and Adults

A basic requirement for many experimental procedures in modern developmental

biology, including live imaging, RNA interference, and gene editing, is the delivery

of synthetic or biological materials into the body of an animal, without disrupting its

health or sacriﬁcing its life. Direct manual injection is one such method and, in the

case of crickets, has been well established and found effective in egg, nymphal,

and adult stages. Two methodological variations are commonly in use for

G. bimaculatus egg injections, diff ering essentially in the number and arrangement

of embryos for injection, and are described in great detail in Horch et al. (2017a) and

Barry et al. (2019). Both methods are thus described below in brief.

The ﬁrst variant of the egg injection method, developed by the Noji lab (University

of Tokushima), involves the construction of a mold to house eggs for injections (Horch

et al. 2017a). Watson chambers (which resemble a rectangular mold) are glued onto

microscopic glass slides using double-sided tape. Embryos are then lined up end to end

along the length of the chamber, using a small stainless-steel spatula. The wall of the

chamber and the adhesive of the double-sided tape (on the slide) help secure and hold

the embryos in place during the injections. The second variant, optimized in the

Extavour lab, uses rectangular troughs made using plastic molds set in low-melting

agarose that hold eggs in place (Kainz et al. 2011; Barry et al. 2019). Using this setup,

over 35 embryos per slide can be prepared for injection simultaneously, making it

efﬁcient in terms of preparation time and the number of embryos injected in one sitting.

Regardless of the egg injection method used, eggs are injected under a dissecting

or compound microscope, using a needle held by a micromanipulator (Fig. 8.6a, b).

The needle must be loaded with the desired injection material and connected to a

pressure source, which may be manual (e.g., a syringe) or electronically controlled

compressed gas (e.g., using a commercial micro-injector). The injected material may

be mixed with a dye that is visible under white light (e.g., phenol red or fast green) or

ﬂuorescent light (e.g., ﬂuorescein- or rhodamine-conjugated dextrans) depending on

the user’s preference. The choice of dye will determine the best microscopy and light

regime to be used for injections. Following injections, embryos are allowed to

develop normally in humid incubators at 28



C on wet paper towels or are sub-

merged in 1 phosphate buffer ed saline in closed petri dishes and monitored daily

until embryos hatch. Eggs of developmental stages that are turgid and under high

pressure, including very early stages in the ﬁrst few hours following fertilization and

middle stages following elongation of the germ band, are more difﬁcult to inject than

earlier stages.

For nymphal and adult injections, a manually held Hamilton syringe or any

automated micromanipulator/microinjector system speciﬁcally designed for delicate

microinjections and capable of injecting nanoliter volumes can be used effectively.

Nymphs and adults are prepared for injections by ﬁrst cooling them on ice to

temporarily immobilize them, and then injected either between the abdominal

segments (A2 and A3) or in the soft tissue between the T3 coxa and the thorax.

For site- or tissue-speciﬁc injections (e.g., the leg or brain), the injection site should

8 The Cricket Gryllus bimaculatus: Techniques for Quantitative .. . 195

Fig. 8.6 Injection and OMMAwell setup. (a) The apparatus used for beveling glass needles for use

in injecting crickets, consisting of a light source, a dissecting microscope, a micromanipulator, and a

beveling stone (Narishige model EG-45 is shown here). (b) Higher magniﬁcation of the beveling

setup shown in (a). (c) Different types of agarose mold inserts used for mounting cricket embryos

for application in OMMAwell or, alternatively, for cricket egg injections. (d) OMMAwell

(Donoughe et al. 2018) schematic for top-loaded microwells, used for injecting and imaging cricket

embryos using a conﬁguration for upright objectives. Different assembly components are shown:

the mold insert (white) consists of wells that will house the cricket embryos and is inverted and

attached to the base of the slide (gray). This is then placed into the upright platform (pink) and

196 A. Kulkarni and C. G. Extavour

be modiﬁed accordingly. However, locales of abundant fat tissue should be avoided

as injection sites in crickets, to help facilitate dispersal of liquid into the body upon

injection, prevent backﬂow of injected mat erial or hemolymph into the needle, and

prevent blockage of ﬁne needle tips with insect tissue. Irrespective of the injection

site, care should be taken while injecting the needle into the nymphal or adult body,

so as to prevent injuring the internal organs, which cou ld potentially kill the animal

or disrupt recovery. Such injuries can be easily avoided by inserting the needle only

deep enough into the animal body to prevent oozing of material at injection.

Other recommendations for successful injections of adult or juvenile

G. bimaculatus include inserting the needle parallel to the body of the insect, rather

than at a perpendicular angle, injecting larger volumes (relative to the insect size) as

multiple pulses of smaller doses rather than all at once, injecting slowly to prevent

leakage, minimizing handling stress for the animal, making sure the needle is not

blocked prior to insertion, and maintaining basic cleanliness and sterility during the

procedure. Following injection, animals should be allowed to recover in isolated cages

with food and water at room temperature before proceeding with the desired study.

8.4.3 High-Throughput Live Imaging of Embryos Using

OMMAwell

Open Modular Mold for Agarose Microwells (OMMAwell) is a simple, reusable,

all-in-one device that allows users to easily mount and simultaneously image dozens

of live G. bimaculatus embryos consistently and economically for 2D and/or 3D

time-lapse analyses of early development (Donoughe et al. 2018). OM MAwell has

the added advantage of being adapta ble and customizable: it has been made to

accommodate the imaging needs of researchers with different experimental designs,

can be used on diverse species (OMMAwell has been successfully designed for and

tested on nine animal species, including many traditional model organisms), and can

be used for both inverted and upright objective microscopes (Fig. 8.6d). With this

device, embryos can be efﬁciently and quickly lined up in arrays of agarose

microwells, whose dimensions and spacing can also be customized as per individual

user needs (Fig. 8.6c) (see Donoughe et al. 2018). In addition, OMMAwell has

reservoirs to hold live imaging media and help maintain specimen-speciﬁc humidity,

osmolarity, and oxygen levels during time-lapse live imaging, thereby enhanci ng

embryonic survival and data quality. This device also allows positional tracking of

⁄

Fig. 8.6 (continued) secured at the desired height with the help of a pin (blue). The assembled

components are then lowered into a petri plate containing molten low-melt agarose and allowed to

set. Following the removal of the mold insert from the cooled and set agarose, specimens are added

into the wells in the petri plate either individually or in bulk and covered with low-melt agarose

(in microliter volumes of up to 100 μl) to hold them in place. Embryos can be oriented carefully

using forceps prior to this step. Once the agarose sets, live-imaging media is poured into the dish

8 The Cricket Gryllus bimaculatus: Techniques for Quantitative .. . 197

individual embryos and permits users to control samp le orientation for imaging. The

OMMAwell microwell array arrangement is also convenient to hold embryos in

place during injections. OMMAwell has been used to image the development of as

many as 102 G. bimaculatus live embryos simultaneously for 12 consecutive days

(Donoughe et al. 2018). Hatching rates of these embryos were not signiﬁcantly

different from the hatching rates of controls, suggesting no lingering effects of

phototoxicity, developmental delays, or defects on these embryos from the use of

OMMAwell.

8.4.4 Gene Expression Analyses Using Embryonic or Whole

Mount In Situ Hybridization

and Immunohistochemistry

The ability to detect mRNAs [using in situ hybridization (ISH)] and proteins

[immunohistochemistry using label ed antibodies (IAb)] of interest is central to

whole mount gene expression analyses in any organism. Standard protocols to

study gene expression in other vertebrate and invertebrate embryos have been

applied successfully in G. bimaculatus (Niwa et al. 2000; Mito and Noji 2008).

These include whole mount in situ hybridizat ion using digoxigenin (DIG)-labeled

antisense RNA probes (as per Wilkinson 1992), protein detection (Patel 1994), and

double in situ hybridization using probes labeled with different haptens (e.g.,

Dietrich et al. 1997). Optimized ISH and IAb protocols have also been developed

in this species for speciﬁc tissues including the brain, nymphal legs, and wings.

Automated medium- or high-throughput gene expression assays on G. bimaculatus

tissues using specialized robots (e.g., Intavis InsituPro VSi) are also possible

(Extavour lab, unpublished).

8.4.5 RNA Interference

The Noji lab pioneered the establishment of RNA interference (RNAi) technology in

the cricket G. bimaculatus (Miyawaki et al. 2004), and many researchers have since

used this technique successfully to deplete mRNAs of multiple target genes in this

species. Four main types of RNAi techniques have been developed for use in crickets:

embryonic, nymphal, parental, and regenerative RNAi (Miyawaki et al. 2004;Mito

et al. 2005;Nakamuraetal.2008a; Mito and Noji 2008; Ronco et al. 2008).

To perform RNAi, double-stranded RNA (dsRNA), preferably 300–500 nucleo-

tides in length, complementary to a region in the G. bimaculatus gene of interest, is

injected into the eggs (embryonic RNAi) or into the body cavity of nymphs

(nymphal RNAi) or adults (paren tal RNAi). Successful concentrations of dsRNA

have been reported to range from 2 to 6 μg/μl (e.g., Kainz et al. 2011). It is

recommended that the dsRNAs designed should match a regio n close to or including

the 3

UTR of the target G. bimaculatus gene, which may minimize off-target effects.

Typical speciﬁcity controls may include testing at least one other dsRNA designed

198 A. Kulkarni and C. G. Extavour

against a nonoverlapping fragment of this same gene. Injecting dsRNA against

exogenous genes not encoded by the cricket genome (e.g., DsRed) and injecting

the buffer alone can also serve as meaningful controls and are thus stro ngly

recommended for every RNAi experiment. Together, these measures can help

researchers distingu ish between speci ﬁc and nonspeciﬁc effects of RNAi, allowing

meaningful interpretation of their results.

RNAi is systemic in G. bimaculatus, such that RNAi-induced phenotypes may be

detected throughout the body of the embryo, nymph, or adult, regardless of the site

of injection. Moreover, the injection of dsRNA into sexually mature adult females

allows for observation of RNAi effects not only in the adult animal itself but also in

its progeny (i.e., eggs) that the animal will lay over the weeks following a

postinjection mating as long as the gene knockdown does not interfere with oogen-

esis, fertilization, or egg laying. Alternatively, nymphal RNAi can be conveniently

used to determine gene functions in postembryonic stages. Regenerative RNAi was

optimized in the Noji lab and has been performed as a speciﬁc application of

nymphal RNAi in the cricket (Nakamura et al. 2008a). For this procedu re, a leg of

a third instar nymph is amputated following dsRNA injection, and the effects of

RNAi are then assessed during the regeneration of the lost leg (which normally

occurs over subsequent molts). Based on these observations, the RNAi response in

crickets can be robust, stable, and even transmissible through subsequent molts

(Nakamura et al. 2008a; Hamada et al. 2015). However, it is recommended that

the robustness of RNAi response, its stability, and duration be determined on a case-

to-case basis; in our hands, there have been instances where the RNAi response for

some genes has lasted only a few days (see Kainz et al. 2011).

8.4.6 Calcium Imaging to Study Neurobiology

and Neuro ethology

The cricket has been an important model for neurobiology and neuroethological

studies, and many physiological techniques are easily applicable to the cricket

(Ogawa and Miller 2017). One such technique is that of calcium imaging, which

uses ﬂorescent dyes and optical methods to monitor the changes in intracellular

levels of calcium ions in live cells and tissues (Neubauer and MacLean 2010),

including cricket neurons. Information on selection of calcium indicators, dye

loading protocols, experimental desig ns, and calcium imaging techniques in the

cricket are well described (Ogawa and Miller 2017). In 2013, Matsumoto and

colleagues successfully expressed Yellow Chameleon (YC) 3.60, a genetically

encoded calcium indicator (GECI) in the cricket brain via electroporation

(Matsumoto et al. 2013), enabling prolonged deep imaging of the cricket brain for

the ﬁrst time. Together with high-resolution microscopy and gene editing tech-

niques, calcium imaging is expected to facilitate major adv ances in our understand-

ing of cricket neurobiology (Ogawa and Miller 2017). Calcium imaging is not

limited to neurobiology, so its successful establishment in the study of cricket

8 The Cricket Gryllus bimaculatus: Techniques for Quantitative .. . 199

nervous tissue suggests that this technique can now also be used to study physiology

in other tissues and cell types in this animal.

8.4.7 High-Sensitivity Trackball Recording Systems

for Studying Phonotaxis and Auditory Neuronal

Plasticity

Acoustic communication is paramount in insects, both within and between species.

Making precise recordings of insect locomotory behavior in response to auditory

stimuli (phonotaxis), such as male calling songs, is often challenging under natural

settings. Various laboratory assays for measuring cricket phonotactic behavior have

been developed, making such studies possible. Examples of such assays include

analyzing the number of crickets that reach an acoustic stimuli or sound target within

adeﬁned time period (Tschuch 1976; Stout et al. 1983), studying cricket behavior in

mazes (Popov and Shuvalov 1977; Rheinlaender and Blätgen 1982), or steering

responses of tethered ﬂying female crickets (Pollack and Hoy 1979). The development

of two different trackball recording systems in crickets has been paramount in

enhancing our understanding of G. bimaculatus auditory steering behavior (Hedwig

2017) and provided detailed insights into insect locomotory behavior in general. All

trackball recording systems measure the movements of the trackball, based on which

insect velocity and direction of insect movement (walking) are inferred, without

allowing the insect to reach the auditory target. In closed-loop trackball systems, the

cricket is allowed to walk and turn freely on the trackball during recordings, with the

trackball compensating for cricket movement by having the ability to counter-rotate.

By contrast, in open-loop systems, the tethered cricket has the ability to walk but not

change its orientation in an acoustic ﬁeld. Due to their sophisticated design, these

trackball recording systems can now easily be integrated into experimental setups

using other forms of recording, including neuro- or electrophysiological and high-

speed video recording experiments. Thus, combined with the GECI YC3.60 discussed

above, and alongside other sophisticated imaging and video recording techniques (see

below), trackball recording systems are expected to provide new insights not only into

cricket biology but also into the study of insect phonotaxis in general.

8.4.8 Automated and Customizable Video Tracking Systems,

Artiﬁcial Crickets, and Cricket Robots for Synthetic

Neuroethology and Social Behavior

Crickets have been used over the past several decades as systems to study behaviors

including mating, ﬂight, aggression, wandering, obstacle avoidance, and importantly,

to study the neurophysiology underlying these processes. When investigating, quan-

tifying, and qualifying animal behavior, dependable and accurate measurement sys-

tems are needed to record animal responses to external stimuli, at both behavioral and

200 A. Kulkarni and C. G. Extavour

physiological levels. The advent of engineering approaches in crickets, especially

robotics, is expected to greatly facilitate such research and is the genesis of the ﬁeld

of cricket synthetic neuroethology. Aonuma and colleagues have described a novel

approach developed for crickets, where provoked animal behavior in response to

computer-generated simulation and robots is captured to effectively bridge the gap

between insect behavior and physiology (Aonuma 2017). Different commercially

available automated video tracking systems designed to follow cricket movement

have also been previously described (Noldus et al. 2001). Recently, another custom-

izable tracking system based on a simple open-source solution called SwisTrack

(Lochmatter et al. 2008) has been introduced for use in crickets. Using this system,

multiple crickets can be video recorded and tracked simultaneously. Because the entire

process is semiautomatic, data collection and its interpretation are more efﬁcient than

previous methods that were based exclusively on manual tracking. Using artiﬁcial

crickets (e.g., Funato et al. 2011; Kawabata et al. 2012;Mizunoetal.2012)orcricket

robots (Funato et al. 2008, 2011) alongside computer modeling is another way of

analyzing cricket behavior that has recently been reported. Further detailed informa-

tion on artiﬁcial crickets, cricket robots, biomimetic robots (Ritzmann et al. 2000), and

behavioral modeling in this species can be found in Aonuma (2017).

8.4.9 Standardized Protocols for Assessing Learning

and Me mory

G. bimaculatus has been reported to have a robust memory and thus has been

exploited for studying the ne ural mechanisms underlying olfactory, auditory, and

visual learning. Mizunami and colleagues have published detailed protocols for

classical conditioning, operant testing, associative learning, memory retention, and

subsequent data analyses in G. bimaculatus (Mizunami and Matsumoto 2017a). A

“classical conditioning and operant testing ” procedure has also been developed in

crickets by these researchers. The establishment of such protocols has resulted in the

elucidation of detailed cellular mechanisms and signaling cascades that are impor-

tant for memory formation in crickets. These studies have additionally revealed that

crickets display unexpected diversity in the mechanisms underlying these proces ses

in compa rison to other insects including Drosophila (Mizunami and Matsumoto

2017b). The use of such classical conditioning paradigms and their variants in

crickets may provide novel breakthroughs in our understanding of learning, cogni-

tion, and memory across animals.

8.4.10 Transgenic Lines

Stable transgenic lines are an invaluable tool for developmental genetics and con-

tribute to the successful establishment of a model animal system. Transgenesis using

P elements, which are the transposon of choice for Drosophila transgenesis (Rubin

8 The Cricket Gryllus bimaculatus: Techniques for Quantitative .. . 201

and Spradling 1982 ), have been found ineffective in crickets, such that other

transposable elements need to be used to achieve transformation in this species.

Zhang and colleagues (2002) showed that Minos transposons (Pavlopoulos et al.

2007) are active in G. bimaculatus embryos and highlighted the possibility of using

these as gene vectors for germ line transformation in this species. However, to our

knowledge, this transposon has not yet been used to establish stable transgenic

cricket lines. Shinmyo et al. (2004) succeeded in somatic transformation of

G. bimaculatus embryos, using the piggyBac transposon (Handler et al. 1998)to

achieve somatic insertion of a construct containing an enhanced green ﬂorescent

protein (eGFP) coding region driven by a G. bimaculatus actin 3/4 promoter.

Construction of plasmids and injection protocols for this line are as described in

Shinmyo et al. (2004) and Zhang et al. ( 2002). Subsequently, this technique has been

optimized to achieve germ line transmission of transgenes (Nakamura et al. 2010).

At the time of writing, a histone2B-GFP (H2B-GFP) transgenic line is stably

maintained in multiple laboratories (Nakamura et al. 2010 ). In this line, the promoter

of the G. bimaculatus actin orthologue (Gb-Actin) drives the expression of the

Gb-histone2B protein tagged with eGFP. This transgene is ubiquitously and consti-

tutively expressed and is maternally contributed to eggs. Based on viability ratios of

the embryos laid by this line, it is likely that the transgene is sublethal in homozy-

gosis (Extavour lab, unpublished observations). As there are no balancer chromo-

somes for G. bimaculatus, heterozygotes must be manually selected at every

generation to maintain the transgene (Extavour lab, unpublished observations).

Zygotic expression of this transgene begins at approximately the fourth day after

egg laying (AEL) at 28



C, and eggs expressing the transgene can then be easily

identiﬁed and selected between 5 and 10 days AEL, based on the presence of

brightly ﬂuorescent nuclei under a ﬂuorescent ster eomicroscope (Fig. 8.2e top and

bottom).

8.4.11 Genome Editing Using CRISPR/Cas9, TALE N,

and Zinc-Finger Nucleases

Sophisticated functional genetics techniques commonly used to modify genomes

in vivo at a speciﬁc site include clustered regularly interspaced palindromic repeats

(CRISPR)/CRISPR- associated nuclease 9 (Cas9), collectively known as the

CRISPR/Cas9 system (Cong et al. 2013), transcription activator-like (TAL) effector

nucleases (TALENs), and z inc-finger nucleases (ZFNs) (Porteus and Carroll 2005;

Moscou and Bogdanove 2009; Remy et al. 2010; Miller et al. 2011; Jinek et al.

2012). All of these approaches work by generating double-stranded breaks in target

DNA sequences, which in turn trigger the cell’s DNA damage response (Remy et al.

2010), and this cellular response can then generate mutations (insertions or dele-

tions) in the targeted gene. All of these techniques are now available and functional

in crickets. The Mito lab established and reported the use of ZFNs and TALENs in

202 A. Kulkarni and C. G. Extavour

crickets in 2012, by successfully creating homozygous genetic knockouts (Watanabe

et al. 2012). CRISPR/Cas9 has also now been effectively used for the generation of

both knock-ins (Horch et al. 2017b) and knockouts (Awata et al. 2015) of cricket

genes. Detailed protocols for knocking-in or knocking-out cricket genes using the

CRISPR/Cas9 method are available in Horch et al. (2017b).

8.4.12 Genomics and Transcriptomics

While no genome sequenc e is currently publicly available for G. bimaculatus,a

number of de novo transcriptomes have been published for this species, providing

gene expression datasets for a number of different speciﬁc tissue types and devel-

opmental stages. To date, these include transcriptomes of the ovaries, embryos, the

prothoracic ganglion, and regenerating legs (Zeng and Extavour 2012; Bando et al.

2013; Zeng et al. 2013; Fisher et al. 2018). Moreover, several transcriptomes

reﬂecting gene expression at different life stages (Berdan et al. 2016), in the male

accessory gland (Andres et al. 2013), fat body and ﬂight muscles of different

ecological morphs (Vellichirammal et al. 2014), and under cold-acclimation condi-

tions (Des Marteaux et al. 2017; Toxopeus et al. 2019), are available for other

species of the genus.

8.5 Novel Insights into Biological Processes Using Forward

Genetics in a Hemimetabolous Insect

G. bimaculatus has been effectively used to study various disciplines of biological

sciences over the past decades. These include early embryonic development and

body patterning, tissue and organ system speciﬁcation, regeneration, body size

regulation, memory and learning, reproductive biology, ecology, physiology, and

endocrinology (reviewed in Horch et al. 2017b). Here, we will therefore refrain from

reiterating the contributions that these resul ts have made to our understanding of

biology. Instead, we will brieﬂy discuss one example of a novel insight that the

biological community has gained through the use of functional genetics in the

cricket.

8.6 The Evolution of the oskar Gene and Its Implications

for Germ-Line Research

Germ cells are the cells that give rise to eggs and sperm. They are therefore an

important cell type in sexually reproducing organisms and are sometimes referred to

as the ultimate totipotent stem cell (Cinalli et al. 2008), because they alone maintain

a genetic link between generations. In a developing embryo, the ﬁrst cells to give rise

8 The Cricket Gryllus bimaculatus: Techniques for Quantitative .. . 203

to the germ cells by clonal mitoti c divisions are known as the primordial germ cells

(PGCs). Across metazoans, PGCs are speci ﬁed using one of two mechanisms

(Extavour and Akam 2003). In some animals, including G. bimaculatus and Mus

musculus inductive cell–cell signaling among neighboring somatic cells instructs

certain cells to adopt PGC fate; this method of PGC speciﬁcation is known as

“induction.” In other animals, including the fruit ﬂy D. melanogaster, the nematode

worm Caenorhabditis elegans, the zebraﬁsh Danio rerio, and the clawed frog

Xenopus laevis, PGCs are instead speciﬁed by “inheritance.” In this mechanism,

PGCs are speciﬁed very early in development through the cytoplasmic inheritance of

a maternally derive d special cytoplasm called “germ plasm,” which often contains

determinants that confer germ-line fate.

oskar is an insect-speciﬁc gene critical for the establishment of germ plasm in

D. melanogaster and is the only gene reported in the animal kingdom to be both

necessary and sufﬁcient for germ cell formation (Lehmann and Nüsslein-Volhard

1986; Ephrussi et al. 1991; Kim-Ha et al. 1991; Ephrussi and Lehmann 1992; Smith

et al. 1992). Following its discovery in D. melanogaster, oskar orthologues were

reported in the genomes of other holometabolous insects known to specify their

germ cells using germ plasm (Goltsev et al. 2004; Juhn and James 2006; Juhn et al.

2008; Lynch et al. 2011). Interestingly, oskar appears to be absent from many insect

genomes that are known to lack germ plasm, including the bee A. mellifera, the

beetle T. castaneum, and the silk moth B. mori (summarized by Quan and Lynch

2016). Based on this observation and the fact that hemimetabolous insects reportedly

lack germ plasm (see Ewen-Campen et al. 2013 and references therein), it was

hypothesized that oskar was a novel gene that arose at the base of the Holometabola,

concurrent with the advent of insect germ plasm (Lynch et al.

2011). However,

Ewen-Campen and colleagues discovered an oskar orthologue in the cricket

G. bimaculatus genome and demonstrated that in this species, oskar is neither

expressed at high levels in PGCs nor required for PGC formation (Ewen-Campen

et al. 2012). Instead, cricket oskar is expressed in the neuroblasts (stem cells that

arise from the neural ectoderm and give rise to the nervous system in Pancrustacea)

of the brain and central nervous system (CNS) of the developing embryo, and

is required for proper embryonic CNS patterning (Ewen-Campen et al. 2012). This

observation, taken together with the reports that D. melanogaster oskar also plays a

neural role (Xu et al. 2013), suggests tw o novel hypotheses: (1) oskar arose at least

50 million years earlier in insect evolution than previously hypothesized, before the

divergence of Hemimetabola and Holometabola, and (2) oskar’s ancestral role in

insects may have been in the nervous system and not in the germ line. This implies

that oskar may have been co-opted for its essential role in holometab olous

germ plasm assembly rather than having originated concurrently with germ plasm

as had been previously sugges ted. This signiﬁcantly changes our understanding of

the evolutionary origins and functional evolution not only of this gene but perhaps

also of insect germ plasm. Moreover, it constitutes an important example of how

novel genes may arise and become co-opted, across evolutionary time scales, to

perform different biological roles in animals.

204 A. Kulkarni and C. G. Extavour

8.7 Genomic Resources in Other Orthopterans

For an organism to become widely used as a research model for comparative or

evolutionary studies, an important contributing factor is whether resources and tools

are also available to study its close relatives. With this in mind, we will discuss

available resources in other orthopterans that may be of use in aiding comparative

work with G. bimaculatus. To our knowledge, at the time of writing, large-scale

genomic resources are available for only two other orthopterans, a locust and a

Hawaiian cricket species.

8.7.1 Resources in Field Crickets

Several transcriptomes are available for tissues and stages of many species of the

genus Gryllus, including (1) gene expression at different life stage s in G. rubens

(Berdan et al. 2016), (2) in the male accessory gland of G. ﬁrmus and

G. pennsylvanicus (Braswell et al. 2006; Andres et al. 2013), (3) fat body and ﬂight

muscles of different ecological morphs of G. ﬁrmus (Vellichirammal et al. 2014),

(4) under cold-acclimation in G. veletis (Des Marteaux et al. 2017; Toxopeus et al.

2019), or (5) adult femur-derived transcriptomes from G. assimilis (Palacios-

Gimenez et al. 2018). Genomic resources are also available for an inbred line of

G. assimilis (Palacios-Gimenez et al. 2018). Outside of the genus Gry llus, large-

scale genomic resources are also available for the Hawaiian cricket Laupala

kohalensis in the form of an EST resource from a nerve cord cDNA library (Danley

et al. 2007) and a de novo draft genome (Blankers et al. 2018). In fact, the

L. kohalensis genome is, to our knowledge, the only published cricket genome to

date. Transcriptomic data are also a vailable for Allonemobius fasciatus embryos

(Reynolds and Hand 2009), male accessory glands of Gryllodes sigillatus (Pauchet

et al. 2015), and the testis, accessory glands, and adult body of Teleogryllus

oceanicus (Bailey et al. 2013).

8.7.2 Resources in Grasshoppers and Locusts

A de novo transcriptome spanning several stages is available for the grasshoppers

Chorthippus biguttulus and Oxya chinensis sinuosa (Kim et al. 2016) and for

nymphs, adult females and males of Xenocatantops brachycerus (Zhao et al.

2018). An organ-speciﬁc transcriptome is available for the gut of Oedaleus asiaticus

(Huang et al. 2017), an EST database exists for transcripts from the central nervous

system of Schistocerca gregaria (Badisco et al. 2011), and a de novo transcriptome

for Tetrix japonica (Qiu et al. 2017) is also available. In addition, other tools

including RNAi have been tested and reported to be successful in S. american a

8 The Cricket Gryllus bimaculatus: Techniques for Quantitative .. . 205

(Dong and Friedrich 2005). Locusta migratoria is a well-studied locust species that

has large-scale genomic resources available in the form of a de novo genome and

transcriptome (Wang et al. 2014) and an EST database from whole body and

dissected organs (Kang et al. 2004; Ma et al. 2006). In addition, RNAi has been

established and reported to be successful in adults, nymphs, and embryos for this

species (He et al. 2006).

8.8 Commercial Importance of Crickets as Edible Insects

and “Food of the Future”

In this chapter we have primarily focused on crickets as emerging evo-devo models.

In this ﬁnal section, we would like to brieﬂy highlight other reasons that crickets are

gaining popularity as study systems. Given their cosmopolitan distribution (all areas

of the world, except the arctic and subarctic regio ns) and over 2400 documented

species, crickets represent the most diverse lineage of “jumping or leaping” insects

(Horch et al. 2017b). While the chirping sounds made by males have historic ally

given them acoustic appeal as pets and in research, many species are now becoming

economically important as an alternative food source for human s (Huis et al. 2013;

Horch et al. 2017b), as feedstock for poultry (Ravindran and Blair 1993), or as ﬁsh

bait (Huis et al. 2013), all of which are multibillion dollar industries. With the human

population predicted to reach nine billion by the year 2050 (Huis et al. 2013), meat

production and consumption is soon expected to reach unsustainable levels (Boland

et al. 2013). Insects, especially crickets, have therefore been proposed and marketed

as a novel, alternative, environmentally efﬁcient food source with high nutritional

value (Oonincx and de Boer 2012 ; Huis et al. 2013; Deroy et al. 2015).

Crickets are reportedly common street snacks in some parts of the world and have

been part of the traditional diet in Thailand, the Lao People’s Democratic Republic,

Vietnam, the Democratic Republic of Congo, and Nigeria for hundreds of years

(Kuhnlein et al. 2009; Huis et al. 2013). As an example, over 20,000 farmers are

reported to rear crickets in Thailand, resulting in an estimated production of over

7500 tons per year in this country alone (Hanboonsong et al. 2013). To increase their

appeal in the West, crickets are now being advertised and marketed to be eaten

whole, in granular or in paste form, and as ingredients in commercially available

ﬂours and protein bars (e.g., Aspire Food Group USA, Inc.). While many species of

crickets are edible (e.g., G. bimaculatus, Gryllodes sigillatus, A. domesticus ,

A. testacea, T. occipitalis, T. mitratus, and Brachytrupes portentosus), to our

knowledge currently only G. bimaculatus, G. sigillatus, and A. domesticus are

farmed economically for human consumption (Huis et al. 2013).

The use of crickets has the potential to change the future of the food industry,

because of how effective they are in maximizing nutrition for minimal resources.

Cricket rearing is comparatively inexpensive, requires a fraction of input resources,

and has fewer negative environmental impacts than rearing traditional vertebrate

206 A. Kulkarni and C. G. Extavour

protein sources (Halloran et al. 2017). Crickets produce only 1% of greenhouse

gases compared to cattle and pigs, in addition to showing an approximate tenfold

reduction in ammonia emission (Oonincx et al. 2010), all relevant factors when

considering sustainable production in the age of clim ate change. They act as a

complete protein source and consi st of over 50% protein by volume (Wang et al.

2004). Other advantages of eating crickets include their high edible weight:

Nakagaki and DeFoliart (1991) have estimated that over 80% of a cricket is edible

and digestible compared to 55% for chicken and pigs and 40% for cattle. This

translates into making crickets twice as efﬁcient as chicken, at least 4 times as

efﬁcient as pigs, and 12 times more efﬁcient as cattle in converting feed into meat

(Huis et al. 2013). As a speciﬁc example, the food conversion efﬁciency of the house

cricket Acheta domesticus has been reported to be ﬁve times higher than beef, and

when their fecundi ty is considered, this has been shown to increase as much as 15- to

20-fold (Horch et al. 2017b; Nakagaki and Defoliart 1991). Farming crickets is

projected to become a multimillion dollar industry, with the US market for edible

insects alone expected to exceed $50 million by as early as 2023 (Ahuja and Deb

2018).

The development of a novel food source like crickets must include assessing the

potential risks involved with consumption of such sources. Therefore, there is

increased interest in understanding the biology of these insects. While studies

addressing entomophagy-induced food allergies (especially ones arising from eating

crickets alone) are few, there is some preliminary evidence to conﬁrm that

crustacean-allergic individuals (or people with seafood allergies) may also show

cross-reactivity to edible insects in general (Srinroch et al. 2015 ; Pener 2016; Ribeiro

et al. 2018). Another study has reported that some individuals can develop asthmatic

symptoms upon ingesting insects belonging to Orthoptera (Auerswald and Lopata

2005). Overall, however, eating and/or exposure to insects is not expected to pose

signiﬁcant risks of allergenic reactions for most people, especially if the individual

has no prior history of arthropod or insect allergen sensiti vity (Huis et al. 2013). In

summary, the disadvantages associated with eating insects like crickets currently

seem few and the advantages many. Consuming reared insects is potentially more

environmentally friendly, nutritious, cheap, and affordable for people in all parts of

the world. Cricket rearing is one way to use land efﬁciently, reduce or lower

pesticide use and greenhouse gas emissions, may boost human and/or animal

immunity (Goodman 1989; Muzzarelli 2010; Taufek et al. 2016), and ﬁnally

improve the livelihood of women and children in rural areas by supporting local

economies (Huis et al. 2013).

8.9 Conclusion

The successful establishment of the many functional and genetic manipulation tools

in crickets has contributed to a new era of non-drosophilid insect research, not

limited to evo-devo research. We hope that scientists from various disciplines feel

8 The Cricket Gryllus bimaculatus: Techniques for Quantitative .. . 207

encouraged to use the cricket as a system to address intriguing questions in their

respective ﬁelds.

Acknowledgments We thank Extavour lab members Aracely Newton and Maitreyi Upadhyay for

helpful comments and Leo Blondel for technical support on the manuscript. This work was

supported by Harvard University.

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