Standardized Approach to Life History Data Collection in Poeciliid Fishes

Abstract

Livebearing fishes in the family Poeciliidae have been essential to testing life history theory. These species are remarkable because males internally inseminate females, and females give birth to free‐swimming young, making these fishes amenable to investigating the evolution of a variety of life history traits, including the timing and nature of maternal reproductive investment, timing of maturity, strategies for maternal provisioning of embryos, and several other classic life history traits. However, researchers vary in the methods that they use to measure these traits, making it difficult to compare findings across studies. Here, I present a standardized approach to studying life history traits in livebearing fishes. I describe methods for preserving samples in the field, for collecting data on a standard set of life history traits, and for processing data in ways that will allow comparisons among studies. I highlight different options in preservation techniques and in data collection that are dependent on the specific questions being addressed. Finally, I argue for a standard approach moving forward to make it possible to complete large‐scale comparative studies to reveal how life history traits have evolved in this important model system.

Full text

1 of 12
Ecology and Evolution, 2025; 15:e70735
https://doi.org/10.1002/ece3.70735
Ecology and Evolution
RESEARCH ARTICLE OPEN ACCESS
Standardized Approach to Life History Data Collection in
Poeciliid Fishes
ErikS.Johnson1,2
1Department of Biology, University of Missouri, St. Louis, Missouri, USA | 2Whitney R. Harris World Ecology Center, St. Louis, Missouri, USA
Correspondence: Erik S. Johnson (smerikjohnson@gmail.com)
Received: 25 July 2024 | Revised: 27 November 2024 | Accepted: 3 December 2024
Keywords: data collection| life history| livebearing fishes| Poeciliidae| standardized approach
ABSTRACT
Livebearing fishes in the family Poeciliidae have been essential to testing life history theory. These species are remarkable be-
cause males internally inseminate females, and females give birth to free- swimming young, making these fishes amenable to
investigating the evolution of a variety of life history traits, including the timing and nature of maternal reproductive investment,
timing of maturity, strategies for maternal provisioning of embryos, and several other classic life history traits. However, re-
searchers vary in the methods that they use to measure these traits, making it difficult to compare findings across studies. Here,
I present a standardized approach to studying life history traits in livebearing fishes. I describe methods for preserving samples
in the field, for collecting data on a standard set of life history traits, and for processing data in ways that will allow comparisons
among studies. I highlight different options in preservation techniques and in data collection that are dependent on the specific
questions being addressed. Finally, I argue for a standard approach moving forward to make it possible to complete large- scale
comparative studies to reveal how life history traits have evolved in this important model system.
1 | Introduction
Ever since Darwin, biologists have tested predictions of evo-
lutionary theory against empirical and experimental findings.
Perhaps no field has enjoyed such a rich blending of theoretical
expectations with empirical data as the study of life history evo-
lution (Roff1993, 2002; Stearns 1992). Indeed, understanding
how organisms adjust their reproductive schedules through-
out their lives in response to different selective agents has pro-
vided powerful insights into the tempo and mode of evolution
in natural systems (Caswell 1983; Gotthard and Nylin 1995;
Nylin and Gotthard 1998; Potter et al. 2021; Salguero- Gómez
and Jones 2017), how evolutionary diversification occurs (Day,
Abrams, and Chase2002; Derrickson and Ricklefs1988; Owens
and Bennett1995; Ricklefs2000; White etal.2013; Winemiller
and Rose 1992), and the extent to which evolution is pre-
dictable and repeatable in the wild (Johnson and Belk 2001;
Moore, Riesch, and Martin 2016; D. N. Reznick, Rodd, and
Cardenas1996; Riesch etal.2014).
Interestingly, much of the progress in understanding life
history evolution has relied heavily on a relatively small
number of model systems for which life history data can be
readily collected, including Arabidopsis plants (Alonso- Blanco
et al. 1999; Donohue 2002; Ellis et al. 2021), Great Tit birds
(Cole etal. 2012; Lack 1954; Wilkin et al. 2006), Drosophila
fruit flies (Schmidt et al. 2005; Schwarzkopf, Blows, and
Caley 1999; Sgrò and Partridge 2000), and others (Berry and
Bronson 1992; Bongers 1999; Harvey, Shorto, and Viney2008;
Svihla1933). Yet, among all model systems, livebearing fishes
in the family Poeciliidae have had a disproportionate impact
as empirical models to test life history theory (Alcaraz and
García- Berthou 2007; Bono, Rios- Cardenas, and Morris 2011;
Downhower, Brown, and Matsui 2000; Evans, Pilastro, and
This is a n open access ar ticle under the terms of t he Creative Commons Attr ibution License, which p ermits use, dis tribution and repro duction in any medium, p rovided the orig inal work is
properly cited.
© 2025 T he Author(s). Ecology and Evolution published b y John Wiley & Sons Lt d.
Erik S . Johnson contribute d to this study.

2 of 12 Ecology and Evolution, 2025
Schlupp 2011; Frías- Alvarez et al. 2014; Golden, Belk, and
Johnson2021; Hulthén etal.2021; Jaime Zúñiga- Vega, Reznick,
and Johnson 2007; Johnson and Belk 2001; José, Rodríguez,
and León 2013; Mukherjee et al. 2014; Pollux et al. 2009;
Reznick, Rodd, and Cardenas 1996; Reznick 1997; Reznick
and Bryga 1996; Riesch, Martin, and Langerhans 2013, 2020;
Riesch et al.2014; Roth- Monzón et al.2021; Schlupp, Taebel-
Hellwig, and Tobler 2010; Weldele, Jaime Zúñiga- Vega, and
Johnson2014; Zúñiga- Vega etal.2024). Pioneering work done in
the Trinidadian guppy (Poecilia reticulata) (Reznick etal.2001;
Reznick and Endler1982; Reznick1997) has inspired dozens of
additional research studies in several other livebearing fishes,
each testing some aspect of life history theory. In fact, we are
fast approaching the point where large- scale comparative stud-
ies across the almost 300 species of Poeciliid fishes are likely to
yield important insights into how reproductive traits evolve in
livebearing organisms.
Given the potential to understand life history evolution in a
large- scale comparative framework, it is critical that the data
evaluated be comparable among species and across different
studies. How life history data have been collected in livebear-
ing fishes has mostly been passed along by word- of- mouth and
by duplicating methods in the literature for the past several
decades, resulting in similarities and also some important dif-
ferences among published studies. Understanding these varied
methodological approaches is important for interpreting past
studies and will be essential for guiding future work. However,
a uniform approach to data collection could ensure more mean-
ingful comparative analyses. To this end, what is needed is a
careful description of various techniques that have been em-
ployed in life history research for livebearing fishes and a stan-
dardized framework for collecting and processing data moving
forward.
Here, I do just that. I provide a brief background on collecting
life history data from livebearing fishes, beginning with how
specimens are collected and preserved in the field. I then de-
scribe a step- by- step technique for processing fish specimens
to collect data for each of several life history traits. I show how
a standard workf low facilitates data collection and preserves
the specimen and its constituent parts for future study. I de-
scribe differences in this process as presented in different re-
search publications, some of which could make it difficult to
compare studies. Finally, I suggest when different approaches
should be deployed for different research questions, and I
argue for a uniform approach to collecting life history data
moving forward so that comparative studies can be effectively
completed.
2 | Life History Data Collection
2.1 | Field Sampling and Specimen Preservation
How specimens are sampled and preserved can have an im-
portant impact on the life history traits that can be evaluated.
Adequate sample sizes are necessary to quantify certain life
history traits, as is sampling across the entire size distribution
of fish in a population. Hence, a carefully designed approach
to both field sampling and specimen preservation techniques is
essential.
2.1.1 | Field Sampling
Selecting sites from which to sample, of course, depends on
the specific research question—previous life history studies in
livebearers have focused on the effects of predation (Jennions
and Telford 2002; Johnson 2001; Johnson and Zúñiga-
Vega 2009; Reznick1982; Reznick, Bryga, and Endler1990),
density (Bassar etal.2013; Johnson and Bagley2011; Reznick
etal.2012), seasonality (Johnson, Tobler, and Johnson2023;
Winemiller1993), and stream gradient (Jaime Zúñiga- Vega,
Reznick, and Johnson2007), all evaluated as agents of natu-
ral selection. Hence, in each of these studies, it is necessary
to sample populations from sites with contrasting selective
regimes. However, regardless of the specific question, all
life history studies should have a common standard for sam-
pling at a particular site. First, it is critical that a sampling
site have enough individuals to characterize the life history.
Unlike other types of studies that require fewer individuals
(e.g., stable isotopes, behavior, and morphometrics), life his-
tory studies require a relatively large sample size. In general,
a minimum of 70 individuals—30 mature females, 20 mature
males, and 20 juveniles—is typically needed to draw robust
conclusions about life history traits within a population.
Mature fish should be sampled from across the size distribu-
tion of individuals within a category. Ideally, fish should be
collected randomly from a particular site to represent the full
range of variation among individuals. It is important to avoid
exhaustive sampling (i.e., repeated seining from one pool or
one reach of a stream) to achieve adequate sample sizes, both
for conservation purposes and to avoid sampling bias within
a collection. Second, when collecting fish from populations
in the wild, it is important to recognize that each site can be
ecologically distinct. Hence, in addition to collecting fish, re-
searchers should gather data on several ecological variables
that can covary with certain life history traits. Quantifying
the density and abundance of aquatic predator species (Gorini-
Pacheco, Zandonà, and Mazzoni2018; Johnson2001; Johnson
and Zúñiga- Vega 2009; Reznick 1982; Reznick, Bryga, and
Endler1990), the presence of potential competitors (Scott and
Johnson2010), t he season in which a sample is taken (Johnson,
Tobler, and Johnson2023; Reznick1989a), estimates of can-
opy cover or resource availability (Grether etal.2001; Walsh
and Reznick 2009), water temperature and water chemistry
(Martin etal.2009; McManus and Travis1998; Riesch, Plath,
and Schlupp 2010), and stream gradient (Jaime Zúñiga- Vega,
Reznick, and Johnson 2007) are all ecological factors that
should be considered when trying to understand variation
among sites in life history traits.
2.1.2 | Specimen Collection and Preservation
Once a fish is collected, it must be preserved so that it can later
be used to provide life history data in the laboratory. Collection
techniques often vary depending on characteristics of the riv-
erine environment, including substrate type, the presence of

3 of 12
boulders or emergent vegetation, the size of the river, etc. Fine-
mesh dip nets and cast nets have both been used in this type
of sampling. However, the most common method to collect
livebearing fishes is the use of a handheld seine. Following col-
lection, live fish can be held in a tank or bucket to ensure that
adequate numbers of specimens will be collected at a particular
site. If not, live fish should be returned to the river. If adequate
numbers can be achieved, specimens should be euthanized and
then fixed in a preservative solution. The ASIH guide for field
collections (Nickum1988) recommends using an overdose of
tricaine methane sulfonate (MS222) or rapid cooling of fish in
ice as appropriate methods to euthanize individuals.
The researcher must choose one of two methods to preserve
specimens: (1) fixation in 100% ethanol or (2) fixation in 40%
buffered formalin. Fish fixed in formalin retain the mass of lipid
material in the specimen, including in the eggs, whereas fish
fixed in ethanol lose their fat content, which is dissolved into the
alcohol solution. Each of these techniques has pros and cons,
and choosing one or the other depends on the research question
asked (as described below). Preserved specimens are then trans-
ported to the laboratory where life history data will be collected.
2.2 | Laboratory Specimen Processing
The first step in processing specimens (Figure1) is to separate
adult males from the remaining specimens. Adult males are dis-
tinguished from juveniles by the presence of a fully developed
gonopodium, the male intromittent organ. In most species, it is
also possible to identify juvenile males by a gonopodium present
that is not fully developed, and in some cases juvenile males can
be identified by the thickening of the third fin ray on the anal
fin (Greven 2011); to see the third fin ray clearly under a mi-
croscope might require removing soft tissue from the surface of
the gonopodium. This initial sorting leaves the researcher with
two groups of male specimens (mature and immature) and a
group that includes both adult females and juveniles (including
both indistinguishable immature males and immature females).
From these three groups of specimens, data on all life history
traits (Box1) for both males and females can be collected. Here,
I detail how these life history trait data are collected—first
FIGUR E  | Standard workflow designed to collect life history data from poeciliid fishes for females (left column) and males (right column). This
workflow spans 2 days to allow for a 24 - h period for specimens to desiccate in a drying oven. The workflow begins on top and progresses down from
step to step. “Data step” refers to each step where data are collected. “Procedure step” refers to any step that is required to manipulate specimens
for data collection or preservation. “Block” refers to a group of steps that are most efficient when performed on all individuals consecutively before
continuing to the next block.
BOX  | Data collected f rom specimens.
All specimens
– Standard length
– Wet mass
Males
– Reproductive state
– Size at maturity
– Gonopodium length
Females
– Reproductive state
– Number of broods
– Number of offspring per brood
– Size of offspring
– Embryo stage
– Brood dry mass
– Somatic dry mass

4 of 12 Ecology and Evolution, 2025
describing data collected from all specimens and then data
uniquely collected from each sex.
2.2.1 | Data to Collect From Each Specimen
The process of data collection is different for males and fe-
males, but several kinds of life history data should be gath-
ered initially from all specimens, including standard length
and wet mass (Box1, details described below). Additionally, if
specimens are used for other analyses such as color, shape, or
functional morphology, photographs should be taken at this
time. Best colors occur in live fish prior to preservation, al-
though in formalin some color remains for a short period of
time following fixation. Photographs for these analyses are
typically taken with a lateral view, with the head to the left to
expose the left flank.
2.2.2 | Male Data Collection Protocol
1. To determine reproductive state, classify males as ma-
ture or immature by examining the anal fin/gonopodium.
A mature male is classified by the presence of a fully de-
veloped gonopodium, which in many species is marked
by the presence of hooks, barbs, serrae, or fleshy palps;
however, gonopodium structure is unique to each species
(Langerhans2011). As a male matures, the appearance of
the gonopodium typically changes from a cloudy appear-
ance to a clear and translucent appearance (Figure 2).
Regardless of structure or appearance, until the gonopo-
dium is completely developed, an individual male cannot
successfully transfer sperm to a female (Greven2011).
2. Use a balance to measure the wet mass of the specimen.
This measure is taken after all excess liquid has been re-
moved from the surface of the fish, typically by using ab-
sorbent tissue, such as ChemWipes or a paper towel.
3. Use calipers to measure the standard length, defined
as the distance from the tip of the snout to the termi-
nus of the vertebral column, readily seen as the point
where the musculature anterior to the caudal fin tapers
(Figure3A).
4. Use calipers to measure the gonopodium length, defined
as the distance from the tip of the gonopodium to the base
of the gonopodium, the point where it extends from the
body (Figure3A).
No additional male life history data are typically collected be-
yond these. However, it is possible to measure the mass of male
gonads (testes) in fresh specimens or specimens that have been
preserved in formalin (Schlupp, Poschadel, and Tobler 2006;
Schrader etal.2012). This is done by dissection, removing the
testes from the body and weighing the gonads in either wet or
dry form (following Schlupp, Poschadel, and Tobler2006 and
Schrader etal. 2012). However, this step is optional given that
life history studies seldom focus on male reproductive invest-
ment—more often, testes mass is measured for studies of sperm
competition (Schrader etal.2012).
FIGUR E  | Images of male gonopodia contrasting juvenile and adult males in four poeciliid fish species: Xenophallus umbratilis (A, B);
Pseudoxiphophorus bimaculatus (C, D); Alfaro cultratus (E, F); Poecilia mexicana (G, H).

5 of 12
2.2.3 | Female Data Collection Protocol
1. Use a balance to measure the wet mass of the specimen.
Prior to weighing, dab the fish dry using a paper towel or
ChemWipes until all excess alcohol is removed from the
sample.
2. Use calipers to measure the standard length of the indi-
vidual (Figure3B).
3. Dissect the female and remove the digestive tract and
embryos (if present). This is done using small dissection
scissors. Make a mid- sagittal cut along the ventral surface
of the fish. Only cut through the body wall and avoid cut-
ting any organs within the body cavity, including the GI
tract. Initiate the cut directly anterior to the insertion of
the anal fin and cut anteriorly to a point directly between
the insertions of the pectoral fins. On the left side of the
fish, beginning where the ventral cut ended, cut dorsally
to a point even with the lateral line (approximately 2/3 of
the way up the body). Make sure to cut under the pectoral
fin to avoid cutting through this structure. Still on the left
side, make a second vertical cut dorsally beginning from
the point of the first incision, and continue up to a point
even with the lateral line (again, approximately 2/3 of the
way up the body). These three cuts should form a flap on
the left side of the fish. Fold this flap upward (dorsally),
exposing the contents of the body cavity. Break the ribs
of the left side by rotating the flap upward using a pair of
forceps—this will open the flap and reveal a window into
the body cavity. Once the window is open and the body
cavity is exposed, identify the digestive tract. Use forceps
to remove the digestive tract at the anus and at the an-
terior end of the body cavity. Preserve the digestive tract
and its contents (if desired) by placing it into a vial filled
with 70% ethanol. Then use forceps to remove embryos
from the body cavity—embryos can often be removed
in a single "sac" composed of the single ovary in these
fishes. However, if the membrane around this sac is bro-
ken, the embryos must be removed one by one. These can
be placed on the specimen plate of the dissecting scope.
After embryos are removed, refold the flap of the female
back into its natural closed position. Place the specimen
into a plastic weigh boat, now ready to be dried.
4. Determine the developmental stage of the em-
bryos by viewing them underneath a dissection scope.
Embryos have historically been staged using either the
Reznick method (Reznick 1981; Reznick, Mateos, and
Springer2002) or the Haynes method (Haynes 1995)—I
compare these two approaches below.
5. Determine the reproductive state of each female spec-
imen. A mature female is classified by the presence of
fertilized, developing embryos, or by the presence of fully
yolked eggs. Females with no embryos or unfertilized em-
bryos are classified as immature.
6. Determine the number of developing broods by ob-
serving whether multiple stages of development are
present among embryos within one specimen (exclud-
ing unfertilized ova). The phenomenon of carrying more
than one brood at different developmental stages simul-
taneously in a single individual is known as superfeta-
tion (Scrimshaw 1944). Fish that carry only one brood
at a time are defined as non- superfetating. In some non-
superfetating species, unfertilized eggs that are yolked
are retained while fertilized embryos develop—these un-
fertilized eggs should not be mistaken for, or counted as,
developing embryos.
7. Count and record the total number of embryos in each
developmental stage. Place these embryos into a plastic
weigh boat.
8. Place both the female and her embryos, both now on
weigh boats, into a drying oven set to 50°C and allow
these specimens to dry for at least 24 h, after which no
additional desiccation occurs. However, drying a spec-
imen for more than 48 h can result in damage to the
specimen.
9. Remove specimens from the drying oven. Immediately
measure the somatic dry mass and brood dry mass
using a balance. It is important to take these measure-
ments immediately, as humidity in the air will cause
specimens to rehydrate (at different rates depending on
the humidity), which is seen as a very minute increase in
weight as the specimen sits on the balance.
10. Finally, rehydrate specimens using distilled or deionized
(DI) water. To rehydrate, place the specimen in a weigh
boat filled with water for approximately 10–20 min, de-
pending on the size of the fish. Adequately rehydrated
samples will be flexible and no longer brittle. At this
FIGUR E  | (A) Image of a male poeciliid, Xenophallus umbratilis,
showing the measurements of standard length and gonopodium length.
(B) Image of a female poeciliid, Xenophallus umbratilis, showing mea-
surement of standard length (see text for description).

6 of 12 Ecology and Evolution, 2025
point, identification tags can be attached using a needle
and museum- grade string. Samples can then be returned
to a jar of 70% ethanol for long- term storage.
After these steps are complete, the researcher will have gathered
all life history trait data that are typically collected in any life
history study.
3 | Life History Data Processing
After collecting the necessary data using the protocols above,
data processing, including some simple calculations, is neces-
sary to quantify some life history traits prior to analysis. I de-
scribe how this data processing is accomplished for the six life
history traits where this is necessary (Box2).
3.1 | Male Size at Maturity
Male size at maturity within populations is calculated by taking
the mean standard length value among all mature males within
the population. Given that males cease growth upon maturation,
the mean body size is frequently used to report size at matu-
rity (Johnson and Bagley 2011). Additionally, some research-
ers choose to report the distribution of male standard lengths,
which can reveal the presence of alternative male mating strat-
egies (Cohen etal.2015; Furness, Hagmayer, etal.2020). It can
also be helpful to examine the size range of maturing males—in
some cases, the size of immature males can overlap with or ex-
ceed the size of mature males (Johnson and Bagley2011).
3.2 | Female Size at Maturity
Female size at maturity within each population is calculated
as follows. Females are typically divided by standard length
into 2 mm length classes. However, smaller size classes might
be preferred (e.g., 1 mm size classes) if the sample size is suffi-
ciently large or if the average body size of the species is particu-
larly small. The size at maturity is then defined as the smallest
length class in which at least half of individuals are mature. I
provide a plot of female dry mass and standard length to visu-
ally illustrate this (Figure4A). Confidence in the size and ma-
turity estimate for females from a population is greatest when
all size classes below the determined size at maturity are com-
posed of immature individuals, and all size classes above the
BOX  | Traits calculated from raw data.
– Male size at maturity
– Female size at maturity
– Maternal resource provisioning strategy (lecithotrophy
or matrotrophy)
– Female reproductive allotment/somatic investment
– Fecundity
– Superfetation
FIGUR E  | (A) Plot showing the relationship between somatic dry mass of females and standard length of females in a population of livebear-
ing fish. Data points represent individual fish, with immature individuals marked in black and mature individuals marked in orange. The 2 mm
size class in which most individuals are mature is marked by a vertical gray bar, in this case the 36- 38 mm size class. Data presented were collected
from Priapichthys annectens at Rio Colorado in Guanacaste, Costa Rica (Johnson, Tobler, and Johnson2023). (B) Plot showing the relationship be-
tween embryo mass and embryo developmental stage (following the Haynes method; see text). The slope of this line is used to describe the maternal
resource provisioning strategy, with positive and flat slopes indicating maternal provisioning throughout embryonic development (i.e., matrotro-
phy) and a negative slope indicating no maternal provisioning after the egg is fertilized (i.e., lecithotrophy). The data presented here come from
Priapichthys annectens taken from a tributary to Rio Cucaracho in Guanacaste, Costa Rica (Johnson, Tobler, and Johnson2023).

7 of 12
determined size at maturity are composed of mature individ-
uals. There may be, however, some exceptions to this pattern
in temperate species that reproduce seasonally (Reznick and
Braun 1987) where mature females that are out- of- season
are not carrying developing embryos. Even in tropical envi-
ronments, some species slow or cease reproduction between
wet and dry seasons (Chapman, Kramer, and Chapman1991;
Chapman and Chapman 1993; Reznick 1989b). These phe-
nomena may bias estimates of female size at maturity. One
indicator that a species is not reproducing year- round is the
presence of females in larger size classes that are found with-
out developing embryos.
3.3 | Maternal Resource Provisioning Strategy
In livebearing fishes, females provision their developing em-
bryos either by loading their eggs entirely with nutrients prior
to fertilization (lecithotrophy) or by provisioning their fertilized
embryos with nutrients throughout embryonic development
(matrotrophy). Most livebearing fishes reproduce year- round;
however, in some species, reproduction can be seasonal, ceas-
ing during the winter or the dry season (Winemiller1993). In
nature, the degree of maternal provisioning can vary along a
gradient from complete lecithotrophy to extreme matrotrophy
(Pollux et al. 2009). This trait is measured by examining the
relationship between the stage of development of embryos (see
below how this is defined) and the mass of an individual em-
bryo. Individual embryo mass is calculated by dividing brood
dry mass by the number of embryos in the brood. Plotting the
relationship between individual embryo mass and developmen-
tal stage reveals if females are provisioning their developing
offspring with nutrients throughout development or prior to
development. A positive slope between these two variables—
showing that embryo mass increases throughout embryonic
development—is evidence for a matrotrophic provisioning strat-
egy. Similarly, a zero slope also demonstrates that females are
provisioning their young throughout development. Only when
the slope is negative, where embryo mass decreases throughout
development, is there evidence for lecithotrophy. To illustrate
this, I provide a plot of individual embryo mass against devel-
opmental stage to visually demonstrate what this looks like for
a lecithotrophic species (Figure4B). The slope of the relation-
ship between embryo mass and developmental stage has been
defined as a "matrotrophy index" (Pollux etal.2009).
3.4 | Female Reproductive Allotment/Somatic
Investment
Reproductive allotment is defined as the total biomass invested
in reproduction, including the mass of all embryos and eggs.
However, because larger females have the capacity for greater
reproductive investment, this trait is often calibrated based on
female body mass (so- called "somatic mass," which is the mass
of the female with both the GI tract and the reproductive tis-
sues and embryos removed). Since reproductive allotment is
often simply represented by the total brood dry mass, female
"somatic mass" of the dried individual is often used as a random
variable in models to control for size variation. Additionally, em-
bryo stage should be included as a random variable as well, as
embryo mass can change across developmental stages. In other
cases, reproductive allotment is measured as a fraction of total
mass (brood dry mass/brood dry mass plus somatic dry mass)—
however, treating this ratio (sometimes called a gonadosomatic
index) as a response variable can be problematic (see below),
and most researchers opt for using somatic mass simply as a
covariate.
3.5 | Superfetation
In some species of livebearing fishes, females can simultane-
ously carry two or more developing broods fertilized at different
times and are therefore at different developmental stages. This
phenomenon is known as superfetation (Scrimshaw1944). The
number of broods in superfetating species can range from two
up to as many as seven (e.g., in the highly superfetating least kil-
lifish, Heterandria formosa; Guzmán- Bárcenas and Uribe2019).
Detecting superfetation requires careful discernment to iden-
tify the presence of embryos at different developmental stages
within a female. It is important to note that in non- superfetating
species, a single brood may appear to have embryos at slightly
different developmental stages, but these are in fact fertilized at
the same time. Additionally, in non- superfetating species, unfer-
tilized eggs can be present in the same tissue as fertilized, devel-
oping embryos. The presence of these unfertilized eggs should
not be interpreted as superfetation.
3.6 | Fecundity
Fecundity is simply the number of offspring produced. In non-
superfetating species, fecundity is measured by counting the
number of embryos that are fertilized and undergoing develop-
ment. In superfetating species, fecundity is usually defined by
the number of embryos in the most developmentally advanced
brood present. Some studies also count the total number of em-
bryos across all broods, regardless of developmental stage. In
some cases, fertilized embryos are spontaneously aborted, a
process known as embryonic regression. These embryos should
not be counted in fecundity measurements. Regressing embryos
are usually found in less advanced stages of development than
the majority of embryos and are characterized by a glassy ap-
pearance, while healthy embryos appear more cloudy (Furness,
Avise, etal.2021; Greven 2011; Norazmi- Lokman, Purser, and
Patil2016).
4 | Toward a Standardized Approach
Although life history data collection in livebearing fishes is
mostly consistent across studies, some variation does exist.
These differences can have an important impact on how some
life history traits are described. Here I highlight some of these
differences, and I explore their consequences.
4.1 | Specimen Preservation Techniques
Two methods of specimen preservation have been used in poe-
ciliids to prepare fish samples for life history study—these are

8 of 12 Ecology and Evolution, 2025
fixation in ethanol and fixation in buffered formalin. There are
advantages and disadvantages to each of these methods, and the
decision to use one or the other is predicated on the nature of the
research question. The choice of preservation technique can af-
fect the measured values of some life history traits. Preservation
in ethanol is often used because it is safer than using formalin
(which is a carcinogen) and more convenient (because long- term
storage of samples is typically in 70% ethanol). Ethanol fixation
also preser ves DNA so that it can be used in future genomic stud-
ies. In contrast, formalin damages DNA, potentially alters nu-
cleotide base pair integrity, and promotes cross- linking among
DNA molecules, making it difficult to sequence DNA (Koshiba
et al. 1993). However, fixing samples in ethanol does impact
some life history traits—ethanol is a solvent for fat. Hence, fix-
ing samples in ethanol results in most fats being removed from
the specimen. Ethanol also dehydrates specimens, which can
alter both internal and external structural traits. This, in turn,
affects measurements of body mass for adult fish and embryos
and of reproductive allotment. In essence, ethanol- fixed fish
provide information on lean mass, with fat content removed. In
contrast, fats are not soluble in formalin. So, an advantage of
formalin- preserved fish is that the fat content of the specimen
remains intact. Most life history studies have used fish fixed in
ethanol, not formalin. Yet, both techniques can be found in the
published literature (Johnson and Bagley 2011). I recommend
preserving specimens in formalin for life history studies and
then retaining a subsample of specimens or tissues for genetic
studies using ethanol preservation. Researchers should be aware
of the effects of different preservation techniques when compar-
ing among collections.
4.2 | Embryo Staging Scales
There are t wo methods of embryo staging for li fe history resea rch:
these are the Reznick method (Reznick1981; Reznick, Mateos,
and Springer2002) and the Haynes method (Haynes1995). Both
approaches assign numbers to embryos to depict their stage of
embryonic development. There are advantages and disadvan-
tages to each method. The Reznick method was developed first
(Reznick1981). This method scores embryos on a scale of 0 to
50, with developmental stages clustered into six major catego-
ries (0 = no development, 10 = uneyed, 20 = ea rly- eyed, 30 = mid-
eyed, 40 = late- eyed, and 50 = very late- eyed). The Reznick
method utilizes increments of 5 to depict broods with individ-
uals spanning two developmental stages (e.g., 35 describes
broods with both mid- eyed and late- eyed embryos). The Haynes
method, developed later, defines 11 developmental stages, with
stages 1 and 2 being small- and medium- sized unfertilized ova,
respectively. Stages 3–11 define a progression of development,
with stage 3 being a fully- yoked egg that is fertilized and stage 11
being a fully developed embryo just prior to parturition. Stages
in between are marked by phenotypic features that can be identi-
fied under a dissecting microscope, with traits such as blastodisc
development, fin development, pigmentation, eye size, and scale
development all featuring in the rubric. In essence, the Haynes
method provides slightly finer resolution in terms of embryonic
traits, although some traits might be difficult to see in preserved
specimens. Unlike the Reznick method, the Haynes method il-
lustrates how developmental stages appear in both lecithotro-
phic and matrotrophic maternal provisioning strategies.
Both the Reznick method and the Haynes method attempt to
categorize stages of embryonic development, but do so by scor-
ing prominent developmental stages that are readily identified
under a dissecting microscope. More detailed staging of em-
bryonic development is possible (Mousavi and Patil 2022), but
it requires freshly harvested embryos from live fish and more
specialized microscopy, which is generally not feasible for field-
collected specimens or from specimens currently housed in
natural history collections. Interestingly, both the Reznick and
Haynes staging methods have been used to plot embryo mass
against developmental stage to score the degree of maternal pro-
visioning, and both methods show a linear relationship between
embryo mass and embryo stage (Reznick and Endler1982). As
described above, this provides a basis for calculating the matrot-
rophy index (Pollux etal.2009). However, a problem with both
methods is that neither approach uses stages that are explicitly
linked to time—that is, neither purports to use stages as a sur-
rogate for developmental age. Indeed, it is likely that embryos
pass relatively rapidly through some stages and spend more
time in others. Unfortunately, such a calibration has not yet
been demonstrated. There may be some advantage to develop-
ing a staging system where each stage is defined by a set period
of time as embryos move through developmental stages, which
would reveal a more accurate view of the pace of development
and which could be useful for comparing among species (E. S.
Johnson, unpublished data). The embryo stage is often used as
a covariate for other life history traits, such as embryo size and
degree of matrotrophy, where it is treated as a continuous vari-
able, when in fact it may not function exactly this way. Although
the Reznick method is perfectly adequate, most life history stud-
ies use the Haynes method, likely due to the graphical distinc-
tion it makes between lecithotrophic and matrotrophic species
(Haynes1995). Given the bias in published studies across spe-
cies toward the Haynes method, this is likely to continue to be
the favored approach. However, a time- based approach that also
depicts distinct stages of embryo development may ultimately be
more informative.
4.3 | Statistical Approaches to Data Processing
Each life history research question can be addressed with a
specific statistical approach appropriate to the particular study.
Prior to these analyses, as I have detailed above, researchers
either directly measure a life history trait (e.g., number of off-
spring, number of broods, male size at maturity, etc.) or they cal-
culate the life history trait (e.g., female size at maturity within a
population, degree of matrotrophy, embryo size, etc.). How these
latter trait values are calculated is important. Because many life
history traits vary with body size, trait values must be size ad-
justed. Some researchers have proposed creating size- adjusted
trait values as response variables prior to conducting statistical
analyses—for example, the gonadosomatic index (GSI) that is
common in fisheries research. However, using ratios (such as
the GSI) can present difficulties in general linear model analy-
ses (Lien, Hu, and Liu2017). That is, how life history traits vary
as a function of size can differ ontogenetically (allometric scal-
ing), and these relationships can also differ among populations
(e.g., Johnson, Tobler, and Johnson2023). Hence, most research-
ers treat covariates as independent factors in statistical models
as part of the analyses. An important caveat in these analyses

9 of 12
is that different life history traits can covary with covariates
in different ways. For example, some work has shown that as
females grow larger, not only do they have more offspring, but
those offspring are smaller (Reznick and Endler1982). In these
cases, it may be necessary to compare life history trait values at
specific female sizes rather than relying on an analysis of cova-
riance where the model assumptions are not met.
5 | Why a Standardized Approach? The Future of
Life History Research in POECILIIDS
The use of poeciliids to test life history theory has an impressive
history (Johnson and Bagley2011). Yet, the stage is set for this
group of fishes to provide new answers to new questions in life
history research, especially questions that examine the evolu-
tion of life history strategies over time and space. Incorporating
a standardized set of techniques to collect life history data will
be critical to facilitate this work.
Comparative studies that examine how life history traits evolve
through evolutionary time offer great promise. Recent phy-
logenetic work shows the relationships among most species
within Poeciliidae (Furness et al. 2019; Rodríguez- Machado
etal.2024), rendering the system particularly useful for com-
parative work. Indeed, some comparative research in poeciliids
has already begun. Small- scale studies comparing a few spe-
cies already exist (Frías- Alvarez etal.2014; Plath etal.2007;
Swenton and Kodric- Brown2012), while genera- wide studies
are less common (but see work done in Poecilia (Pires and
Reznick 2018), Limia (Cohen et al. 2015), and Phallichthys
(Regus et al.2013)). Family- wide life history studies are few
but appear to be very promising. For example, research on vivi-
parity in livebearing fishes comparing multiple species across
the entire family has revealed insights into the evolution of
placentation (Furness etal.2019; Furness, Avise, et al.2021;
Meredith etal.2010), superfetation (Furness et al. 2019), and
matrotrophy (Olivera- Tlahuel etal.2015). These types of stud-
ies are particularly adept at exploring if the presence of cer-
tain life history traits primes the evolution of subsequent traits
(e.g., matrotrophy preceding superfetation). Family- wide com-
parative studies might also yield additional insights into other
aspects of life history evolution, including ontogenetic shifts
in patterns of reproductive allotment, understanding how sex-
ual conflict affects life histories, and basic insights into how
male life histories evolve. Finally, comparisons among spe-
cies across the phylogeny can also reveal evolutionary inter-
actions between life history strategies and other traits known
to vary among livebearing fishes, including alternative male
mating strategies and functional morphological adaptations.
While these interactions have been probed within some spe-
cies (Domínguez- Castanedo et al. 2023; Furness, Hagmayer,
etal. 2021), understanding how life history strategies evolve
over time, and in relation to the evolution of other traits, is
likely only to be uncovered within a phylogenetic framework
when multiple species are compared.
Ultimately, addressing any of these questions will require hav-
ing comparable life history data for multiple species. Hence,
adopting a standardized approach to collect, manipulate, and
analyze data to score life history traits in livebearing fishes will
allow this model system to continue to contribute to our under-
standing of life history theory now and into the future.
Author Contributions
Erik S. Johnson: conceptualization (equal), data curation (equal), for-
mal analysis (equal), funding acquisition (equal), investigation (equal),
methodology (equal), project administration (equal), resources (equal),
software (equal), supervision (equal), validation (equal), visualization
(equal), writing – original draft (equal), writing – review and editing
(equa l).
Acknowledgments
I am grateful to several scientists who have directly (with hands on
demonstration) or indirectly (through published work) shared with me
their approaches to studying Poeciliid fish life histories. Publishing
costs were paid for by the Erik S. Johnson fund for open access fees.
David Reznick and an anonymous reviewer provided helpful feedback
on this manuscript.
Conflicts of Interest
The author declares no conflicts of interest.
Data Availability Statement
All data presented in this paper are available either in the paper itself or
in other referenced papers.
References
Alcaraz, C., and E. García- Berthou. 2007. “Life History Variation of
Invasive Mos quitofish (Gambusia holbrooki) Along a Salinity Gr adient.”
Biological Conservation 139, no. 1–2: 83–92.
Alonso- Blanco, C., H. Blankestijn- de Vries, C. J. Hanhart, and M.
Koornneef . 1999. “Natural Allelic Var iation at Seed Size Loci i n Relation
to Other Life History Traits of Arabidopsis thaliana.” Proceedings of the
National Academy of Sciences of the United States of America 96, no. 8:
4710– 4717.
Bassar, R. D., A. Lopez- Sepulcre, D. N. Reznick, and J. Travis. 2013.
“Experimental Evidence for Density- Dependent Regulation and
Selection on Trinidadian Guppy Life Histories.” American Naturalist
181, no. 1: 25–38.
Berry, R. J., and F. H. Bronson. 1992. “Life History and Bioeconomy of
the House Mouse.” Biological Reviews of the Cambridge Philosophical
Society 67, no. 4: 519–550.
Bongers, T. 1999. “The Maturity Index, the Evolution of Nematode Life
History Traits, Adaptive Radiation and Cp- Scaling.” Plant and Soil 212,
no. 1: 13–22.
Bono, L. M., O. R ios- C ardenas, and M. R. Morr is. 2011. “Alternat ive Life
Histories in Xiphophorus multilineatus: Evidence for Different Ages at
Sexual Maturity and Growth Responses in the Wild.” Journal of Fish
Biology 78, no. 5: 1311–1322.
Caswell, H. 1983. “Phenotypic Plasticity in Life- History Traits:
Demographic Effects and Evolutionary Consequences.” American
Zoologist 23, no. 1: 35–46.
Chapman, L. J., and C. A. Chapman. 1993. “Desiccation, Flooding,
and the Behavior of Poecilia gillii (Pisces: Poeciliidae).” Ichthyological
Exploration of Freshwaters 4, no. 3: 279–287.
Chapman, L. J., D. L. Kramer, and C. A. Chapman. 1991. “Population
Dynamics of the Fish Poecilia gillii (Poeciliidae) in Pools of an
Intermittent Tropical Stream.” Journal of Animal Ecology 60, no. 2:
441– 453.

10 of 12 Ecology and Evolution, 2025
Cohen, S. N., J. U. Regus, Y. Reynoso, T. Mastro, and D. N. Reznick.
2015. “Comparative Life Histories of Fishes in the Subgenus Limia
(Pisces: Poeciliidae).” Journal of Fish Biology 87, no. 1: 100–114.
Cole, E. F., J. Morand- Ferron, A. E. Hinks, and J. L. Quinn. 2012.
“Cognitive Ability Influences Reproductive Life History Variation in
the Wild.” Current Biology: CB 22, no. 19: 1808–1812.
Day, T., P. A. Abrams, and J. M. Chase. 2002. “The Role of Size- Specific
Predation in the Evolution and Diversification of Prey Life Histories.”
Evolut ion; International Journal of Organic Evolut ion 56, no. 5: 877–887.
Derrickson, E. M., and R. E. Ricklefs. 1988. “Taxon- Dependent
Diversif ication of Life- History Traits and the Perception of Phylogenetic
Constraints.” Functional Ecology 2, no. 3: 417.
Domínguez- Castanedo, O., M. Suárez- Rodríguez, I. Solano- Zavaleta,
and J. J. Zúñiga- Vega. 2023. “Do Male Reproductive Traits Evolve at the
Intraspecific Level in Response to the Amount of Placentotrophy in a
Genus of Viviparous Fishes?” Evolutionary Ecology 37: 927–943.
Donohue, K. 20 02. “Germination T iming Inf luences Natura l Selection on
Life- History Characters in Arabidopsis thaliana.” Ecology 83, no. 4: 1006.
Downhower, J. F., L. P. Brown, and M. L. Matsui. 2000. “Life History
Variation in Female Gambusia hubbsi.” Environmental Biology of Fishes
59, no. 4: 415–428.
Ellis, T. J., F. M. Postma, C. G. Oakley, and J. Ågren. 2021. “Life- History
Trade- Offs and the Genetic Basis of Fitness in Arabidopsis thaliana.”
Molecular Ecology 30, no. 12: 2846–2858.
Evans, J. P., A. Pilastro, and I. Schlupp. 2011. Ecolog y and Evolution of
Poeciliid Fishes. Chicago, IL, USA: University of Chicago Press.
Frías- Alvarez, P., C. Macías Garcia, L. F. Vázquez- Vega, and J. J.
Zúñiga- Vega. 2014. “Spatial and Temporal Variation in Superfoetation
and Related Life History Traits of Two Viviparous Fishes: Poeciliopsis
gracilis and P. infans.” Science of Nature 101, no. 12: 1085–1098.
Furness, A. I., J. C. Avise, B. J. A. Pollu x, Y. Reynoso, and D. N. Reznick.
2021. “The Evolution of the Placenta in Poeciliid Fishes.” Current
Biology: CB 31, no. 9: 2004–2011.e5.
Furness, A. I., A. Hagmayer, and B. J. A. Pollux. 2020. “Size- Dependent
Male Mating Tactics and Their Morphological Correlates in Poecilia
gillii.” Biological Journal of the Linnean Society. Linnean Society of
London 131, no. 4: 880–897.
Furness, A. I., A. Hagmayer, and B. J. A. Pollux. 2021. “Female
Reproductive Mode Shapes Allometric Scaling of Male Traits in Live-
Bearing Fishes (Family Poeciliidae).” Journal of Evolutionary Biology
34, no. 7: 1144–1155.
Furness, A. I., B. J. A. Pollux, R. W. Meredith, M. S. Springer, and D.
N. Reznick. 2019. “How Conflict Shapes Evolution in Poeciliid Fishes.”
Nature Communications 10, no. 1: 3335.
Golden, K. B., M. C. Belk, and J. B. Johnson. 2021. “Predator
Environment Does Not Predict Life History in the Morphologically
Constrained Fish Alfaro cultratus (Cyprinodontiformes: Poeciliidae).”
Frontiers in Ecolog y and Evolution 9: 1–11.
Gorini- Pacheco, B., E. Zandonà, and R. Mazzoni. 2018. “Predation
Effects on Matrotrophy, Superfetation and Other Life History Traits in
Phalloceros harpagos.” Ecology of Freshwater Fish 27, no. 1: 442–452.
Gotthard, K., and S. Nylin. 1995. “Adaptive Plasticity and Plasticity as
an Adaptation: A Selective Review of Plasticity in Animal Morphology
and Life History.” Oikos 74, no. 1: 3–17.
Grether, G. F., D. F. Millie, M. J. Bryant, D. N. Reznick, and W. Mayea.
2001. “Rain Forest Canopy Cover, Resource Availability, and Life
History Evolution in Guppies.” Ecology 82, no. 6: 1546–1559.
Greven, H. 2 011. “Gonads, G enitals, and Reproductiv e Biology.” Ecology
and Evolution of Poeciliid Fishes 3: 3 –17.
Guzmán- Bárcenas, M. G., and M. C. Uribe. 2019. “Superfetation in
the Viviparous Fish Heterandria formosa (Po eci liidae).” Journal of
Morphology 280, no. 5: 756–770.
Harvey, S. C., A. Shorto, and M. E. Viney. 2008. “Quantitative Genetic
Analysis of Life- History Traits of Caenorhabditis elegans in Stressful
Environments.” BMC Evolutionary Biology 8: 15.
Haynes, J. L. 1995. “Standardized Classification of Poeciliid
Development for Life- History Studies.” Copeia 1: 147–154.
Hulthén, K., J. S. Hill, M. R. Jenkins, and R. B. Langerhans. 2021.
“Predation and Resource Availability Interact to Drive Life- History
Evolution in an Adaptive Radiation of Livebearing Fish.” Frontiers in
Ecology and Evolution 9: 1–14.
Jaime Zúñiga- Vega, J., D. N. Reznick, and J. B. Johnson. 2007. “Habitat
Predicts Reproductive Superfetation and Body Shape in the Livebearing
Fish Poeciliopsis turrubarensis.” Oikos (Copenhagen, Denmark) 116, no.
6: 995–1005.
Jennions, M., and S. Telford. 2002. “Life- History Phenotypes in
Populations of Brachyrhaphis episcopi (Poeciliidae) With Different
Predator Communities.” Oecologia 132, no. 1: 44–50.
Johnson, E. S., M. Tobler, and J. B. Johnson. 2023. “Seasonality
Predicts Variation in Life History Phenotypes in the Livebearing Fish
Priapichthys annectens.” Biological Journal of the Linnean Society.
Linnean Society of London 142: blad119.
Johnson, J. B. 2 001. “Adaptive Li fe- History Evolution in the Li vebearing
Fish Brachyrhaphis rhabdophora: Genetic Basis for Parallel Divergence
in Age and Size at Maturity and a Test of Predator- Induced Plasticity.”
In Evolution; International Journal of Organic Evolution 55, no. 7:
1486–1491.
Johnson, J. B., a nd J. C. Bagley. 2011. “Ecological Dr ivers of Life- History
Divergence.” In Ecology and Evolution of Poeciliid Fishes, 38–49.
London, UK: University of Chicago Press.
Johnson, J. B., and M. C. Belk. 2001. “Predation Environment Predicts
Divergent Life- History Phenotypes Among Populations of the
Livebearing Fish Brachyrhaphis rhabdophora.” Oecologia 126, no. 1:
142–149.
Johnson, J. B., and J. J. Zúñiga- Vega. 2009. “Differential Mortality
Drives Life- History Evolution and Population Dynamics in the Fish
Brachyrhaphis rhabdophora.” Ecology 90, no. 8: 2243–2252.
José, L., R. Rodríguez, and G. León. 2013. “Life- History Patterns of
Cuban Poeciliid Fishes.” Zoo Biology 32: 251–256.
Koshiba, M., K. Ogawa, S. Hamazaki, T. Sugiyama, O. Ogawa, and
T. Kitajima. 1993. “The Effect of Formalin Fixation on DNA and the
Extraction of High- Molecular- Weight DNA From Fixed and Embedded
Tissues.” Pathology, Research and Practice 189, no. 1: 66–72.
Lack, D. 1954. “The Natural Regulation of Animal Numbers.” ht tps ://
doi. org/ 10. 2307/ 4511784.
Langerhans, R. B. 2011. “Genital Evolution.” In Ecology and Evolution
of Poeciliid Fishes, 228–242. London, UK: University of Chicago Press.
Lien, D., Y. Hu, and L. Liu. 2017. “A Note on Using Ratio Variables in
Regression Analysis.” Economics Letters 150: 114–117.
Martin, S. B., A. T. Hitch, K. M. Purcell, P. L. Klerks, and P. L. Leberg.
2009. “Life History Variation Along a Salinity Gradient in Coastal
Marshes.” Aquatic Biology 8: 15–28.
McManus, M. G., and J. Travis. 1998. “Effects of Temperature and
Salinity on the Life History of the Sailfin Molly (Pisces: Poeciliidae):
Lipid Storage and Reproductive Allocation.” Oecologia 114, no. 3: 317–
325. https:// doi. org/ 10. 1007/ s0044 20050453.
Meredith, R. W., M. N. Pires, D. N. Reznick, and M. S. Springer.
2010. “Molecular Phylogenetic Relationships and the Evolution of the
Placenta in Poe cilia (Micropoec ilia) (Poeciliidae: Cy prinodontiformes).”
Molecular Phylogenetics and Evolution 55, no. 2: 631–639.

11 of 12
Moore, M. P., R. Riesch, and R. A. Martin. 2016. “The Predictability and
Magnitude of L ife- History Divergenc e to Ecological Agents of S election: A
Meta- Analysis in Livebearing Fishes.” Ecology Letters 19, no. 4: 435–442.
Mousavi, S. E., and J. G. Patil. 2022. “Stages of Embryonic Development
in the Live- Bearing Fish, Gambusia holbrooki.” Developmental
Dynamics: An Official Publication of the American Association of the
Anatomists 251, no. 2: 287–320.
Mukherjee, S., M. R. Heithaus, J. C. Trexler, J. Ray- Mukherjee, and J.
Vaudo. 2014. “Perceived Risk of Predation Affects Reproductive Life-
History Traits in Gambusia holbrooki, but Not in Heterandria formosa.”
PLoS One 9, no. 2: e88832.
Nickum, J. G. 1988. “Full issue article volume 13, issue 2.” Fisheries 13,
no. 2: 1–62.
Norazmi- Lokman, N. H., G. J. Purser, and J. G. Patil. 2016. “Gravid
Spot Predicts Developmental Progress and Reproductive Output in a
Livebearing Fish, Gambusia holbrooki.” PLoS One 11, no. 1: e0147711.
Nylin, S., and K. Gotthard. 1998. “Plasticity in Life- History Traits.”
Annual Review of Entomology 43: 63–83.
Olivera- Tlahuel, C., A. G. Ossip- Klein, H. S. Espinosa- Pérez, and J. J.
Zúñiga- Vega. 2015. “Have Superfetation and Matrotrophy Facilitated the
Evolution of Larger Offspring in Poeciliid Fishes?” Biological Journal of
the Linnean Society Linnean Societ y of London 116, no. 4: 787–804.
Owens, I. P. F., and P. M. Bennett. 1995. “Ancient Ecological
Diversification Explains Life- History Variation Among Living Birds.”
Proceedings. Biological sciences 261, no. 1361: 227–232.
Pires, M. N., and D. N. Reznick. 2018. “Life- History Evolution in the
Fish Genus Poecilia (Poeciliidae: Cyprinodontiformes: Subgenus
Pamphorichthys): An Evolutionary Origin of Extensive Matrotrophy
Decoupled From Superfetation.” Biological Journal of the Linnean
Society. Linnean Society of London 125, no. 3: 547–560.
Plath, M., A. M. Makowicz, I. Schlupp, and M. Tobler. 2007. “Sexual
Harassment in Live- Bearing Fishes (Poeciliidae): Comparing Courting
and Noncourting Species.” Behavioral Ecolog y: Off icial Journal of the
International Society for Behavioral Ecology 18, no. 4: 680–688.
Pollux, B. J. A., M. N. Pires, A. I. Banet, and D. N. Reznick. 2009.
“Evolution of Placentas in the Fish Family Poeciliidae: An Empirical
Study of Macroevolution.” Annual Review of Ecology, Evolution, and
Systematics 40: 271.
Potter, T., R. D. Bassar, P. Bentzen, etal. 2021. “Environmental Change,
if Unaccounted, Prevents Detection of Cryptic Evolution in a Wild
Population.” American Naturalist 197, no. 1: 29–46.
Regus, J. U., J. B. Johnson, S. A. Webb, and D. N. Reznick. 2013.
“Comparative L ife Histories of Fishes i n the Genus Phallichthys (Pisces:
Poec iliidae).” Journal of Fish Biology 83, no. 1: 144–155.
Reznick, D. 1981. “‘Gr andfather Eff ects’: The Genetics of Inter population
Differences in Offspring Size in the Mosquito Fish.” Evolution;
International Journal of Organic Evolution 35, no. 5: 941–953.
Reznick, D. 1982. “The Impact of Predation on Life- History Evolution
in Trinidadian Guppies—Genetic- Basis of Observed Life- History
Patterns.” Evolution; International Journal of Organic E volution 36, no.
6: 1236–1250.
Reznick, D., G. Buckwalter, J. Groff, and D. Elder. 2001. “The Evolution
of Senescence in Natural Populations of Guppies (Poecilia reticulata):
A Comparative Approach.” Experimental Gerontology 36, no. 4–6:
791–812.
Reznick, D., and J. A. Endler. 1982. “The Impact of Predation on
Life History Evolution in Trinidadian Guppies (Poecilia reticulata).”
Evolution; International Journal of Organic E volution 36, no. 1: 160–177.
Reznick, D. A., H. Bryga, and J. A. Endler. 1990. “Experimentally
Induced Life- History Evolution in a Natural Population.” Nature 346,
no. 6282: 357–359.
Reznick, D. N. 1989a. “Life- History Evolution in Guppies: 2.
Repeatability of Field Observations and the Effects of Season on Life
Histories.” Evolution 43, no. 6: 1285. https:// doi. org/ 10. 2307/ 2409363.
Reznick, D. N. 1989b. “L ife- History Evolution in Guppies: 2. R epeatability
of Held Observations and the Effects of Season on Life Histories.”
Evolut ion; Internati onal Journal of Organi c Evolution 43, no. 6: 12 85–1297.
Reznick, D. N. 1997. “Life History Evolution in Guppies (Poecilia re-
ticulata): Guppies as a Model for Studying the Evolutionary Biology of
Aging.” Experimental Gerontology 32, no. 3: 245–258.
Reznick, D. N., R. D. Bassar, J. Travis, and F. Helen Rodd. 2012. “Life-
History Evolution in Guppies VIII: The Demographics of Density
Regulation in Guppies (Poecilia reticulata).” Evolution; International
Journal of Organic Evolution 66, no. 9: 2903–2915.
Reznick, D. N., and B. Braun. 1987. “Fat Cycling in the Mosquitofish
(Gambusia affinis): Fat Storage as a Reproductive Adaptation.”
Oecologia 73, no. 3: 401–413.
Reznick, D. N., and H. A. Bryga. 1996. “Life- History Evolution in
Guppies (Poecilia reticulata: Poeciliidae). V. Genetic Basis of Paral lelism
in Life Histories.” American Naturalist 147, no. 3: 339–359.
Reznick, D. N., M. Mateos, and M. S. Springer. 2002. “Independent
Origins and Rapid Evolution of the Placenta in the Fish Genus
Poeciliopsis.” Science 298, no. 5595: 1018–1020.
Reznick, D. N., F. H. Rodd, and M. Cardenas. 1996. “Life- History
Evolution in Guppies (Poecilia reticulata: Poeciliidae). IV. Parallelism
in Life- History Phenotypes.” American Naturalist 147, no. 3: 319–338.
Ricklefs, R. E. 2000. “Density Dependence, Evolutionary Optimization,
and the Diversi fication of Avian Life Histories.” Condor 102, no. 1: 9 –22.
Riesch, R., R. A. Martin, and R. B. Langerhans. 2013. “Predation's Role
in Life- H istory Evolution of a Livebear ing Fish and a Test of the Trexler-
DeAngelis Model of Maternal Provisioning.” American Naturalist 181,
no. 1: 78–93.
Riesch, R., R. A. Martin, and R. B. Langerhans. 2020. “Multiple Traits
and Multifarious Environments: Integrated Divergence of Morphology
and Life History.” Oikos (Copenhagen, Denmark) 129, no. 4: 480–492.
Riesch, R., M. Plath, and I. Schlupp. 2010. “Toxic Hydrogen Sulfide and
Dark Caves: Life- History Adaptations in a Livebearing Fish (Poecilia
mexicana, Poeciliidae).” Ecology 91, no. 5: 1494–1505.
Riesch, R., M. Plath, I. Schlupp, M. Tobler, and R. Brian Langerhans.
2014. “Colonisation of Toxic Environments Drives Predictable Life-
History Evolution in Livebearing Fishes (Poeciliidae).” Ecology Letters
17, no. 1: 65–71.
Rodríguez- Machado, S., D. J. Elías, C. D. McMahan, A. Gruszkiewicz-
Tolli, K. R. Piller, and P. Chakrabarty. 2024. “Disentangling Historical
Relationships With in Poeciliidae ( Teleostei: Cy prinodontiformes) Using
Ultraconserved Elements.” Molecular Phylogenetics and Evolution 190:
107965.
Roff, D. A., ed. 1993. Evolution of Life Histories. New York, NY, USA:
Chapman and Hall.
Roff, D. A. 2 002. Life Histor y Evolution. Sunderland, MA, USA: Sinauer.
Roth- Monzón, A. J., M. C. Belk, J. J. Zúñiga- Vega, and J. B. Johnson.
2021. “What Drives Life- History Variation in the Livebearing Fish
Poeciliopsis prolifica? An Assessment of Multiple Putative Selective
Agents.” Frontiers in Ecology and Evolution 8: 1–12.
Salguero- Gómez, R., and O. Jones. 2017. “Life History Trade- Offs
Modulate the Speed of Senescence.” In The Evolution Of Senescence in
the Tree of Life, 403–421. Cambridge, UK: Cambridge University Press.
Schlupp, I., J. Poschadel, and M. Tobler. 2006. “Male size polymorphism
and testis weight in two species of mollies (Poecilia latipinna, P. me xi-
cana, Poeciliidae, Teleostei).” Zeitschrift Für Fischkunde 8: 9–16.

12 of 12 Ecology and Evolution, 2025
Schlupp, I., A. Taeb el- Hellwig, and M. Tobler. 2010. “Equal Fecu ndity in
Asexual and Sexual Mollies (Poecilia).” Environmental Biology of Fishes
88, no. 2: 201–206.
Schmidt, P. S., L. Matzkin, M. Ippolito, and W. F. Eanes. 2005.
“Geographic Variation in Diapause Incidence, Life- History Traits,
and Climatic Adaptation in Drosophila melanogaster.” Evolution;
International Journal of Organic Evolution 59, no. 8: 1721–1732.
Schrader, M., J. J. Apodaca, P. S. D. Macrae, and J. Travis. 2012.
“Population Density Does Not Inf luence Male Gonadal Investment in
the Least Killifish, Heterandria formosa.” Ecology and Evolution 2, no.
12: 2935–2942.
Schwarzkopf, L., M. W. Blows, and M. J. Caley. 1999. “Life- History
Consequences of Divergent Selection on Egg Size in Drosophila melano-
gaster.” American Naturalist 154, no. 3: 333–340.
Scott, L. E., and J. B. Johnson. 2010. “Does Sympatry Predict Life
History and Morphological Diversification in the Mexican Livebearing
Fish Poeciliopsis baenschi?” Biological Journal of the Linnean Society
Linnean Society of London 100, no. 3: 608–618.
Scrimshaw, N. S. 1944. “ Superfetation in Poecil iid Fishes.” Copeia 1944 ,
no. 3: 180.
Sgrò, C. M., and L. Partridge. 2000. “Evolutionary Responses of the
Life History of Wild- Caught Drosophila melanogaster to Two Standard
Methods of Laboratory Culture.” American Naturalist 156, no. 4: 341.
Stearns, S. C. 1992. The Evolution of Life Histories. New York, NY, USA:
Oxford University Press.
Svihla, A. 1933. “A Comparative Life History Study of the Mice of the
Genus Peromyscus.” Murrelet 14, no. 2: 46.
Swenton, D. M., and A. Kodric- Brown. 2012. “Habitat and Life History
Differences Between Two Species of Gambusia.” Environmental Biology
of Fishes 94, no. 4: 669–680.
Walsh, M. R., and D. N. Reznick. 2009. “Phenotypic Diversification
Across an Environmental Gradient: A Role for Predators and Resource
Availability on the Evolution of Life Histories.” Evolution; International
Journal of Organic Evolution 63, no. 12: 3201–3213.
Weldele, M. L., J. Jaime Zúñiga- Vega, and J. B. Johnson. 2014. “Life
History of Gambusia vittata (Pisces: Poeciliidae).” Southwestern
Naturalist 59, no. 4: 449–460.
White, A. E., Y. J. Li, V. Griskevicius, S. L. Neuberg, and D. T. Kenrick.
2013. “Putting all Your Eggs in One Basket: Life- History Strategies, Bet
Hedging, and Diversification.” Psychological Science 24, no. 5: 715–722.
Wilkin, T. A., D. Garant, A . G. Gosler, and B. C. Sheldon. 2006. “Density
Effec ts on Life- History Tra its in a Wild Popul ation of the Great Tit Parus
major: Analyses of Long- Term Data With GIS Techniques.” Journal of
Animal Ecology 75, no. 2: 604–615.
Winemiller, K. O. 1993. “Seasonality of Reproduction by Liverbearing
Fishes in Tropical Rainforest Streams.” Oecologia 95, no. 2: 266–276.
Winemiller, K. O., and K. A. Rose. 1992. “Patterns of Life- History
Diversification in North American Fishes: Implications for Population
Regulation.” Canadian Journal of Fi sheries and Aquatic Sciences 49, no.
10: 2196–2218.
Zúñiga- Vega, J. J., C. Olivera- Tlahuel, N. Saleh- Subaie, and M. Suárez-
Rodríguez. 2024. “Life Histories of Poeciliid Fishes: Searching for a
Size- Dependent Trade- Off Between Number and Size of Offspring.”
Ichthyological Research 71, no. 1: 106–118.