APU Oceanography The Ecological Conditions Questions

Assignment Steps1.
Read the “Guide to Reading Academic Research Papers” located in the
Canvas assignment folder.
2. In the file attached there is a journal article under name (Dolphins & Sponges)
folder to read, annotate, and analyze.
3. Read and annotate the paper using the steps outlined in the guide. Do not
just highlight text. Make notes about the paper, write any questions, identify
and define words you did not understand, and comment on parts of the
paper you like/dislike or agree/disagree with. You may do this by hand or
digitally.
4. Critically analyze one of the figures included in the paper. Include the figure in
a word document and answers to the following questions.
•
•
•
1. What is the purpose of this figure? How do you know?
2. What key elements are included in this figure that allows you to understand
what is going on? What information does the author include that helps the
reader understand the figure?
3. Do you think this is the best way to represent this data visually? Why? If
not, how could the information be conveyed more clearly (different types of
figures, breaking the figure into multiple parts, a table, etc.)?
1. Write a one-page summary of the paper that answers the questions below:
•
•
•
•
1. What was the experimental goal? Why was it important to the broader
world?
2. Briefly explain the methods of this paper. If you do not understand the
methods, explain what parts you had difficulty with.
3. What were the main conclusions of the paper? How do they connect to the
broader world and previous studies? What are some of the possible
implications of this experiment? Do you agree with the conclusions that the
author drew?
4. What did you like and dislike about this paper? What would you change,
and what is one takeaway you can use in your writing?
Your summary should not exceed one page and should include a proper citation and
reference
for the paper. For this assignment, you can discuss your paper as a group. However,
each person must submit their own files. For grading, you should submit three files:
1. A copy of the paper with your annotations. If you choose to print the paper
and do the annotations by hand, please scan the paper and create a pdf file. I
recommend an app called Genius Scan for this; however, there are many
similar options. If you need help with this, please contact one of us.
2. A one-page summary of the paper that answers the questions above and
includes the proper citation and reference. For the citation and reference, you
may use the formatting of whatever scientific journal is most relevant to your
field of study.
3. A single page with the figure you choose to analyze and the answers to the
questions above.
The Ecological Conditions That Favor Tool Use and
Innovation in Wild Bottlenose Dolphins (Tursiops sp.)
Eric M. Patterson*, Janet Mann
Department of Biology, Georgetown University, Washington, D.C., United States of America
Abstract
Dolphins are well known for their exquisite echolocation abilities, which enable them to detect and discriminate prey
species and even locate buried prey. While these skills are widely used during foraging, some dolphins use tools to locate
and extract prey. In the only known case of tool use in free-ranging cetaceans, a subset of bottlenose dolphins (Tursiops sp.)
in Shark Bay, Western Australia habitually employs marine basket sponge tools to locate and ferret prey from the seafloor.
While it is clear that sponges protect dolphins’ rostra while searching for prey, it is still not known why dolphins probe the
substrate at all instead of merely echolocating for buried prey as documented at other sites. By ‘sponge foraging’ ourselves,
we show that these dolphins target prey that both lack swimbladders and burrow in a rubble-littered substrate. Delphinid
echolocation and vision are critical for hunting but less effective on such prey. Consequently, if dolphins are to access this
burrowing, swimbladderless prey, they must probe the seafloor and in turn benefit from using protective sponges. We
suggest that these tools have allowed sponge foraging dolphins to exploit an empty niche inaccessible to their non-toolusing counterparts. Our study identifies the underlying ecological basis of dolphin tool use and strengthens our
understanding of the conditions that favor tool use and innovation in the wild.
Citation: Patterson EM, Mann J (2011) The Ecological Conditions That Favor Tool Use and Innovation in Wild Bottlenose Dolphins (Tursiops sp.). PLoS ONE 6(7):
e22243. doi:10.1371/journal.pone.0022243
Editor: Sarah Frances Brosnan, Georgia State University, United States of America
Received April 23, 2011; Accepted June 17, 2011; Published July 20, 2011
Copyright: ß 2011 Patterson, Mann. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: Support for this study was provided by The National Geographic Society Young Explorers Grant (http://www.nationalgeographic.com/field/grantsprograms/young-explorers/) and The Explorers Club Exploration Fund Grant (http://www.explorers.org/index.php/expeditions/funding/expedition_grants) to
EMP and the National Science Foundation (http://www.nsf.gov/) 0918303, 0847922, 0316800 to JM. Field gear was supplied at discounted rates by Airline Supply
by J. Sink (http://www.airlinebyjsink.com/) and Splash Dive Center of Alexandria, VA (http://www.splashdivecenter.com/). The orthophoto in Figure 2 was
supported by NSF grant OCE0526065 to Dr. Mike Heithaus and DEC. The funders had no role in study design, data collection and analysis, decision to publish, or
preparation of the manuscript.
Competing Interests: The authors have the following competing interest: Field gear was supplied at discounted rates by Airline Supply by J. Sink (http://www.
airlinebyjsink.com/) and Splash Dive Center of Alexandria, VA (http://www.splashdivecenter.com/). There are no patents, products in development or marketed
products to declare. This does not alter the authors’ adherence to all the PLoS ONE policies on sharing data and materials, as detailed online in the guide for
authors.
* E-mail: emp46@georgetown.edu
(Figure 1B) as they probe the rough seafloor (Figure 1C) while
searching for hidden prey (Figure 1D). Once prey have been
extracted, dolphins drop their sponges, occasionally surface for a
quick breath, chase and consume their prey, and finally, return to
pick up their sponges and continue foraging [13,14]. Spongers are
suspected to target one or few benthic prey species, including the
barred sandperch, Parapercis nebulosa, previously mis-identified as
Parapercis clathrata [14], whose confamilials are consumed by
bottlenose dolphins elsewhere [17]. The sponge is thought to
function as a shield by providing protection from the sharp and
rough seafloor, and possibly venomous or spiny benthic marine
organisms, while dolphins search for and extract prey [13,14].
Dolphins use a single sponge for an average of 68647 (SD)
minutes (Max = 4.4 hrs, Min = 3 minutes, N = 125 sponging
bouts) before dropping it to search for a replacement presumably
because the sponge has lost its protective value. However, why
dolphins continuously probe the substrate when searching for prey
is unclear given that at other locations (e.g. crater feeding in the
Bahamas [12]) dolphins detect buried prey indirectly via
echolocation and minimize contact with the seafloor until prey
are located. In fact, delphinids’ target detection ability using
echolocation is quite impressive [18,19] and has long been used by
the U.S. Navy to locate buried mines [20]. So in contrast to other
Introduction
Tool use [1,2] has long been of interest to biologists,
anthropologists, and psychologists because of its role in cognition,
culture, and hominid evolution [3–5]. Studying tool use in animals
provides insight into the social, ecological, and evolutionary
contexts in which it arises [6]. In mammals and birds, tool use
positively correlates with brain size, social transmission, and
innovation [7] and is considered to be a sign of cognitive capacity,
i.e., problem solving ([8,9] but see [10,11]). Most animal tools are
used during foraging, especially extractive foraging [1,12]. In
Shark Bay, Western Australia some bottlenose dolphins (Tursiops
sp.) use marine basket sponge tools to protect their rostra during
foraging [13–16]. Thus far, we know that sponge foraging,
hereafter sponging, is primarily a female behavior, appears to be
vertically socially transmitted [14], and is limited to 54 animals
(hereafter the spongers) in the eastern gulf of Shark Bay [14]. This
solitary behavior occurs in deep channels, requires long dives, and
consumes the majority of spongers’ activity budgets, yet does not
appear to have any fitness costs [14].
On several occasions, during exceptional visibility, researchers
have directly observed dolphins wearing marine sponges they have
removed from the substrate (Figure 1A) over their rostra
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Wild Dolphin Tool Use
Figure 1. Sponging in Shark Bay. (A) marine basket sponge (Echinodyctium mesenterinum), (B) dolphin wearing a sponge on its rostrum, (C)
substrate littered with rock, shell, and debris, (D) hiding prey, barred sandperch (Parapercis nebulosa). All photographs taken by Eric M. Patterson.
doi:10.1371/journal.pone.0022243.g001
extractive tool users [12], dolphins appear to have the anatomical
machinery necessary for the task at hand leaving one to consider:
Why do Shark Bay dolphins probe or skim the substrate with their
rostra and risk injury instead of simply echolocating for prey?
It is well known that the major acoustic backscatter of fishes
(over 90%) comes from the gas-filled swimbladder [21–30]. This
is not surprising as sound waves most readily echo when
encountering a difference in medium density (i.e. liquid to gas)
[31]. Fishes without swimbladders have relatively weak acoustic
signals, as fish flesh has an acoustic impedance only 10% greater
than water [29]. Many fish species have lost their swimbladders,
presumably as an adaptation to their benthic or deep-sea lifestyle
[32]. While some odontocetes are capable of echolocating
swimbladderless prey (cephalopod detection by Globicephala,
Ziphiidae, and Physeteridae [33,34]), the majority of these
cetaceans’ prey are free swimming, not buried [35], and
echolocation in these cephalopod specialists appears to be
modified by longer click intervals and higher source levels when
compared to bottlenose dolphins [36]. We hypothesized that
spongers probe the substrate because they target prey that lack
swimbladders and thus are difficult to detect with echolocation.
Moreover, when these prey are at least partially buried beneath
a debris-laden substrate, which causes interfering reverberation
and echo clutter (echoes from objects other than the targeted
prey) [37], the effectiveness of echolocation is reduced even
further. In contrast, dolphins that crater-feed in the Bahamas
[38] appear to target buried prey with swimbladders [39] in an
uncluttered, soft sand substrate that is less likely to injure
dolphins’ rostra or dramatically interfere with echolocation [40].
While some echolocation has been documented during sponging,
it may only be useful once prey have been extracted and
dolphins have dropped their sponges since the sponge itself is
likely to interfere with echolocation by obstructing the sound
receiving lower jaw and the sound emitting melon [41]. Thus, we
predicted that the majority of prey that spongers encounter lack
swimbladders.
PLoS ONE | www.plosone.org
Results
Of the 134 prey extracted during 13.3 hours of human
Sponging (Video S1) on both transect and verification dives
(Figure 2), 78% lacked swimbladders (Table 1). In contrast only
19% of prey from all Non-Sponging dives lacked swimbladders
(Table 1). Barred sandperch (Figure 1D), which lack swimblad-
Figure 2. Sponging Map. Boat launch site (Monkey Mia), dolphin
sponge foraging sightings, transects, and verification dive sites in Shark
Bay, Western Australia.
doi:10.1371/journal.pone.0022243.g002
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Table 1. Prey abundance from Sponging and Non-Sponging, pooled from both transects and verification dives.
Common Name
Family
Species (if known)
Sponging Abundance
Non-Sponging Abundance
Swimbladder
barred sandperch
Pinguipedidae
Parapercis nebulosa
87
15
N[53]*
sand lizardfish
Synodontidae
Synodus dermatogenys
9
0
N[52]*
cuttlefishes
Sepiidae
5
0
N+
stingrays
Dasyatidae
1
1
N+
lefteye flounders
Bothidae
1
0
N[32]*
painted maskray
Dasyatidae
Neotrygon leylandi
1
0
N+
tasselsnout flathead
Platycephalidae
Thysanophrys cirronasa
1
0
N[52]*
purple tuskfish
Labridae
Choerodon cephalotes
23
23
Y[52,54]*
freckled goatfish
Mullidae
Upeneus tragula
4
0
Y[52]
wrasses
Labridae
2
0
Y[52,54]*
striped whiptail
Nemipteridae
Pentapodus vitta
0
29
Y*
margined coralfish
Chaetodontidae
Chelmon marginalis
0
6
Y[52]#
blackspot tuskfish
Labridae
Choerodon schoenleinii
0
5
Y[52,54]*
bluntheaded wrasse
Labridae
Thalassoma amblycephalum
0
2
Y[52,54]*
humpback batfish
Ephippidae
Platax batavianus
0
2
Y[52]#
yellowtail clownfish
Pomacentridae
Amphiprion clarkii
puffers
Tetraodontidae
0
1
Y[52]#
0
1
Y[62]*
Numbers represent reference(s) used to determine swimbladder status.
*Dissected in this study, +Swimbladder well known to be absent in entire family,
#
Additional family encountered on verification dives.
doi:10.1371/journal.pone.0022243.t001
ders, were by far the most common prey extracted during
Sponging, constituting 65% of the total count, but only made up
18% of Non-Sponging prey. Purple tuskfish (Choerodon cephalotes),
which possess swimbladders, were second in abundance during
both Sponging and Non-Sponging at 17% and 27% respectively;
however, 74% of purple tuskfish extracted during Sponging were
from locations where less sponging has been documented
(verification dives, Figure 2), and this species was only extracted
when divers probed small seagrass tufts which are both
uncharacteristic of channel habitat and less likely to harm
dolphins’ rostra. Striped whiptail (Pentapodus vitta), which possess
swimbladders, were the predominant prey during Non-Sponging at
34%, but not extracted at all during Sponging. No additional
families were extracted during Sponging on verification dives,
although verification dives did yield 3 additional Non-Sponging
families (Table 1), indicating that our transects are representative
of spongers’ prey, but not of all non-burrowing prey in the
eastern gulf of Shark Bay, which is not surprising.
The ratio of prey without swimbladders to those with
swimbladders was significantly higher during Sponging compared
to Non-Sponging on video transects (Figure 3, Wilcoxon signedrank test W = 28, P = 0.016), demonstrating that dolphins
encounter a greater proportion of swimbladderless prey when
sponging than is available to them without disturbing the
substrate. Furthermore, the abundance of prey extracted during
Sponging on transects was significantly greater than that for the
same prey families during Non-Sponging on transects (Figure 4,
Wilcoxon signed-rank test W = 28, P = 0.016), indicating that
these prey are primarily concealed in the substrate and that
sponging is an effective method of extraction. Together these
results show that sponging dolphins extract concealed swimbladderless prey, and do so with greater efficiency than could be done
without a sponge tool. Finally, a permutation test revealed that
the number of prey families extracted during Sponging that lack
PLoS ONE | www.plosone.org
swimbladders was significantly greater than expected when
compared to 27 years of data from the Shark Bay Dolphin
Research Project’s long-term study (P = 0.0132, Table S1),
suggesting that sponging is the primary way dolphins access
swimbladderless prey.
Figure 3. Ratio of prey without swimbladders (SB) to prey with
swimbladders during both Sponging and Non-Sponging on
transects. Data were transformed (+1) before ratios were calculated
to correct for undefined ratios in samples with zero individuals in either
group. Wilcoxon Sign Rank Test W = 28, *P = 0.016.
doi:10.1371/journal.pone.0022243.g003
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Wild Dolphin Tool Use
breathe before diving back down in pursuit, as has been regularly
documented during our long-term study [14]. New photographs
from 2010 of a sponger consuming a small red and brown fish
provided further evidence that spongers prey on barred sandperch.
Sponging dolphins may gain several benefits from targeting these
prey. First, barred sandperch exhibit consistent, predictable behavior
enabling dolphins to employ a single stereotypic sponging behavior. If
dolphins extracted a variety of prey species, all having different antipredator tactics, a uniform sponging behavior would not be as
effective. Second, similar to some foods extracted by primate tool
users [12], barred sandperch are reliable and can easily and
frequently be extracted with a sponge, one every nine minutes
during human Sponging. However, the average barred sandperch
collected was small, only 12.664.7 (SD) cm in length (Max = 23 cm,
Min = 6 cm, N = 21), which may explain why spongers are more
specialized and dedicate more time to foraging than other dolphins in
Shark Bay [14]. Finally, extracted foods are often high energy,
premium foods [12]. Since fishes with decreased swimbladder
volumes typically have increased lipid content [27], the barred
sandperch may provide sponging dolphins with an energy-rich meal,
similar to some insect larvae extracted by tool-using birds [47].
Accordingly, several barred sandperch have been stored for
nutritional analysis. Thus, while requiring more effort than freeswimming prey, barred sandperch likely provide these tool users with
a small, yet predictable, reliable, and possibly energy rich food source.
This highly specific tool use has implications for cognition and
brain evolution among cetaceans and could even be considered a
case of problem solving, a phenomenon difficult to document in
the wild, but well established in studies of captive bottlenose
dolphins [48]. Our study demonstrates how bottlenose dolphins
might use these skills in their natural environment and provides
insight into the ecological and evolutionary pressures that promote
higher-level cognition. Spongers may have solved the problem of
detecting and extracting swimbladderless prey from below a sharp
and rough substrate by probing the seafloor with a soft sponge
tool. This solution appears to have been adopted at least twice, as
unrelated dolphins residing 110 km away in the western gulf of
Shark Bay also sponge forage [15]. While this tool use requires
sophisticated object manipulation, it appears to provide spongers
with equal fitness compared to the rest the population [14].
Due to the difficulty of observing marine fauna, most studies of tool
use focus on terrestrial organisms. Using novel underwater techniques, we show that sponge tool-using dolphins target buried prey
that lack swimbladders, particularly barred sandperch. Such prey are
difficult to detect with echolocation [21–30], which, when paired with
Shark Bay’s cluttered channel substrate, explains why dolphins probe
the seafloor and benefit from using sponge tools. Similar to ant-fishing
chimpanzees whose tool use is a function of prey type [49], dolphin
tool use directly relates to the physical characteristics of their prey. As
such, this study emphasizes the critical role ecological factors play in
explaining behavioral complexity.
Figure 4. Abundance of prey species extracted during Sponging
and abundance of these same families during Non-Sponging on
transects. Wilcoxon sign-rank test W = 28, *P = 0.016.
doi:10.1371/journal.pone.0022243.g004
Discussion
Our results demonstrate that sponging dolphins regularly
encounter swimbladderless prey that are concealed beneath a
rubble-littered substrate. Fish are comprised primarily of water (fish
flesh: 82% water, 17% protein, and 0.35% fat by weight [42]) so the
density of fish (less the swimbladder) is only slightly greater than
seawater (1.076 compared to 1.026 g/cm3 [43]). As such,
swimbladderless prey have little acoustic impedance and are
difficult to detect with echolocation [21–30]. To make matters
more difficult, Shark Bay channels are strewn with fragmented rock,
shell, and coral that are not only likely to injure dolphins’ rostra, but
also create an echo-cluttering environment leaving dolphins
‘acoustically blind’ to swimbladderless prey. Similarly, echolocating
bats seem to have trouble detecting prey on the surface of ponds that
are covered with duckweed [44]. Our data demonstrate that
spongers have developed a way of effectively extracting hidden prey
by probing the substrate with protective sponge tools. Furthermore,
when compared to the rest of the Shark Bay population, sponging
dolphins appear to specialize in prey that lack swimbladders
allowing them to occupy an empty ecological niche.
Alternatively, it is possible that dolphins could listen for
soniferous benthic prey and simply use sponges for extraction. In
fact, several species of echolocating bats passively listen for preygenerated sounds to detect insects in highly cluttered environments
[45]. However, we believe this is unlikely for Shark Bay dolphins
since only two prey families that were infrequently extracted
during human Sponging are reported to be soniferous [46], and
both possess swimbladders making them detectable with echolocation anyway. The remaining swimbladderless prey are unlikely
to be soniferous since the primary sonic mechanism in fishes is
swimbladder movement [46]. Furthermore, fishes mainly produce
sound for intraspecific communication [46], not while hidden or
buried in the substrate as observed in this study.
The predominant prey extracted during human Sponging was the
barred sandperch whose behavior was strikingly consistent with
dolphin sponging behavior [14]. When barred sandperch were
disturbed during human Sponging, they swam a few meters away
and returned to the substrate often without reburying. This would
give dolphins time to drop their sponges and quickly surface to
PLoS ONE | www.plosone.org
Materials and Methods
Ethics Statement
All animal work was approved by the Georgetown University
Animal Care and Use Committee (GUACUC) under permits 07041 and 10-023. Observational and field studies were approved by
the Department of Environment and Conservation of Western
Australia (DEC) under permits SF007418 and SF006897.
Study Site
The Shark Bay Dolphin Research Project has an extensive 27year database that includes demographic, genetic, association, life4
July 2011 | Volume 6 | Issue 7 | e22243
Wild Dolphin Tool Use
history, ecological, and behavioral data on .1,400 dolphins (past
and present) residential to a 300 km2 area (25u479S, 113u439E).
Habitat consists of embayment plains (5–13 m), shallow sand flats
(0.5–4 m), seagrass beds (0.5–4 m), and bisecting deep channels
(7–13 m). One area in particular, just north of Monkey Mia
(Figure 2), is extremely well sampled because it is very close to our
boat-launching site. As such, spatial patterns of dolphin foraging
behavior in this location are unlikely the result of a bias in
sampling effort. Using historical data, dolphin-sighting locations
were projected using ArcGIS Map 9.3 (WGS 1984 UTM Zone
49S) to determine locations where dolphins sponge forage. Seven
semi-permanent transects were established using 1 m long star
picket metal posts with attached buoys in locations with the highest
numbers of recorded sponging observations. Each 100 m transect
was ,200 m from adjacent transects and further split into two
portions (NW and SE) by a mid-stake. The NW portion of each
transect was dedicated to systematic observational-video sampling,
while the SE portion was designated for prey sample collection.
encounter. Swimbladder status for prey families not collected
and prey from video only identified to the level of family was
determined using primary literature [32,52–62]. All data in
Table 1 and Table S1 follow the currently accepted scientific and
common names according to Froese and Pauly (2008) [63] and all
analyses were performed at the family level.
Data processing and statistical analysis
Many of the species encountered were quite small, averaging
less than 7 cm in length. Such small prey are unlikely to be
targeted during sponging because prey this size can easily be
obtained at the surface in all habitats, even by young calves
[64,65]. Thus, these prey were removed from prey abundance
data. A total of 19 prey encountered could not be identified to the
level of family; however, all were estimated to be less than 7 cm in
length and thus excluded from the final data set. There is a chance
prey were missed during video logging or were simply not
captured on film; however, it is likely that all such prey would also
be less than 7 cm in length and thus excluded since prey larger
than this would be obvious to divers and not overlooked.
The ratio of prey without swimbladders to those with swimbladders
was compared between Sponging and Non-Sponging on transects using a
Wilcoxon signed-rank test. Here, prey abundance data were
transformed by adding one to all samples before ratios were
calculated to correct for samples with zero individuals in either
group, which results in an undefined ratio. We also compared the
abundance of prey families extracted during Sponging on transects to
the abundance of these same families during Non-Sponging on transects
using a Wilcoxon signed-rank test. Finally, we compared data from
Sponging dives to data from our long-term study. For this final analysis
we used a two-tailed permutation test to compare the observed
number of families with and without swimbladders from Sponging to
the expected number based on combined Sponging and historical prey
data. We re-sampled (with replacement) 8 prey families 10,000 times
from 29 possible families (Table S1) and determined the likelihood of
obtaining our observed human Sponging data by chance. While Table 1
present abundance data pooled from all dives for descriptive purposes,
all statistical analyses were performed only using the systematic
transect data for which we could be sure that the substrate traversed
during Sponging and Non-Sponging were equal. All statistical tests were
performed in R 2.12.1 statistical environment (R Development Core
Team, 2011) and considered significant for P,0.05.
Data Collection
For all 7 transects, two certified divers swam out a 50 m tape
measure well above the substrate to connect the NW stake to the
mid-stake for initial transect setup. After waiting several minutes,
both divers then swam back towards the NW stake along one side
of the transect line near the substrate and filmed a ,2 m wide
belt transect [50,51] to determine prey availability near the
seafloor without disturbing the substrate (Non-Sponging). Next,
divers swam back along the other side of the transect (,2 m to
the side of the tape measure) towards the mid-stake with one
diver pushing a 2 m long pole with a dead marine sponge
attached along the substrate to ferret prey in the same manner as
seen by sponging dolphins (Sponging), and the other diver filming
this human Sponging with a Sony HDR-XR500V HD video
camera in an AquaticaHD housing (Video S1). All dives were
performed on an Airline Supply R360XL Hookah System by J.
Sink, and were swam at a consistent speed of ,17 m min21. On
the NW portion of each transect, all prey were simply filmed and
allowed to swim away. However, on the SE side of the transects
several sample specimens of all species encountered (except
Dasyatidae and Sepiidae which all lack swimbladders) were
collected using hand nets for identification and dissection.
Transect sampling was performed on two different occasions
for repeatability, but replicates were averaged to form a single
transect value. To confirm that our fine scale study in this well
sampled area was representative of greater patterns in the bay, in
particular to be sure we had extracted all possible Sponging prey
species, we also performed verification dives in all other general
locations where sponging has been observed (Figure 2). On these
verification dives no tape measure was laid, but divers performed
and filmed both Non-Sponging and Sponging as described above. If
any new species were encountered, sample specimens were
collected for identification and dissection. Only one infrequent
prey species was too fast for divers to catch, Upeneus tragula, but
other species in the same genus are known to have swimbladders
[52]. Historical prey species were gathered from the long-term
Shark Bay Dolphin Research Project’s database and combined
with families extracted during Sponging to create a total of 29
possible prey families. Families were used instead of individuals
due to the potential biases in observing accurate quantities of
species consumed (e.g. researchers observe dolphins consuming
surface dwelling prey more often than benthic prey since dolphins
regularly consume these prey near the surface, in plain site).
Using families allows us to avoid these potential biases and
explore how dolphins consume the richness of prey they
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Supporting Information
Table S1 Sponging and historical prey families. Numbers
represent reference(s) used to determine swimbladder status.
*Dissected in this study, +Swimbladder well known to be absent
in entire family.
(DOCX)
Video S1 Divers performing human Sponging. Prey in order of
appearance: Sepia sp., Parapercis nebulosa, Parapercis nebulosa, Neotrygon
leylandi, Parapercis nebulosa, Parapercis nebulosa, Parapercis nebulosa,
Synodus dermatogenys.
(MP4)
Acknowledgments
We thank our field assistants James Barnao, Anne Cotter, and Jacob
Swartz, and all members of the Mann lab, especially Ewa Krzyszczyk, for
providing new photos of a sponger with its prey. We thank all past and
present members of the Shark Bay Dolphin Research Project who
contributed to the long-term data. Western Australian Museum’s Dr. Barry
Hutchins and Ms. Sue Morrison helped with prey identification and
provided advice on field sampling methods, while Dr. Jane Fromont
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Wild Dolphin Tool Use
identified sponge species. We also thank the Shark Bay Ecosystem
Research Project for help in the field, and the Monkey Mia Dolphin Resort
and the Department of Environment and Conservation of Western
Australia (DEC) for field and logistical support.
Author Contributions
Conceived and designed the experiments: EMP. Performed the experiments: EMP JM. Analyzed the data: EMP. Contributed reagents/
materials/analysis tools: EMP. Wrote the paper: EMP JM. Collected most
of the data on sponging in Figure 2: JM.
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Published in
Towards Data Science
https://towardsdatascience.com/guide-to-reading-academic-research-papers-c69c21619de6
Kyle M Shannon
Jul 21, 2018
Guide to Reading Academic Research Papers
Learn to tackle this laborious process with a systematic approach!
My desk is full of papers. Slightly shuffled around for dramatic purposes of course.
Working in data science and machine learning is an exciting and challenging field. New
techniques and tools are constantly percolating and honestly, it can feel overwhelming. Many of
these new developments are found and first revealed in academic research articles. Extracting
knowledge from these articles is difficult because the intended audience of these papers tend to
be other researchers. Yet in order to stay current reading papers is an essential skill — luckily
one that can be improved with diligence and practice.
In graduate school, you get good (should get good…) at reading papers and ingesting research.
Not everyone will get training in this skill, that doesn’t mean you shouldn’t benefit from the
knowledge these papers hold. Public tax money is how most of this research gets funded
anyways! The goal here is to democratize academia, just a bit, and provide you with a
scaffolding to apply when walking through a paper.
The way I read papers is not significantly unique, but it is effective and has served me well.
Keep in mind it is not the only way, many techniques exist and as you read more and more I am
sure you will find your own unique style.
This guide is broken down as follows:
● Learning this skill will help you! I promise
● So I hear reading a paper is difficult. Why?
● How are papers typically organized?
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My “bullet proof” approach to reading papers
Tools to help you get the job done
Why Learn to Read Papers?
Reading papers certainly builds character because It often takes many hours and there is no
guarantee you walk away with the whole story. This is not to disparage you, but merely to be
open and transparent. Reading papers is difficult, there are no two ways about it. Advances in
fields such as machine learning, deep learning, data science, databases, and data engineering
often come in the form of academic research, whose language is that of academic papers.
Think about some of the techniques you might use: Convolutional Neural Networks, PCA, and
AdaBoost (even Deep Boosting!). These all came out of research, and yes they all have papers.
Also, consider that there are many papers on the application and use of these techniques and
when you are trying to solve a specific problem, these papers can be critical. Beyond staying
current with research it is also worth traveling to the past and giving older papers a read. You
will learn so much. I promise.
Looking at the field of deep learning it seems as though a new critical paper is coming out every
few days or weeks. The only way to stay on top of it is to get a hold of the paper and give it a
read.
Where the Difficulties Arise…
Here is a figure from a 2017 scientific paper¹ by Hubbard and Dunbar, about reading scientific
papers. Scientific Paper inception!
Fig 2. Different sections of scientific papers are considered easy to read and important at
different stages of academic careers.
A: The proportion of participants considering a section easy to read (presented as ‘Somewhat
easy’, ‘easy’ ‘very easy’ combined) as a function of career stage. Results of Chi-square tests are
indicated on the left hand side. B: The mean importance rank of sections as a function of career
stage. Error bars are omitted from individual points for clarity, with the sole error bar in grey
representing the largest 95% confidence interval for any of the data points. Asterisks above data
points indicate significant differences in response compared with the previous career stage as
determined by Mann-Whitney post-hoc tests.
One unsurprising result indicates the further an academic progresses into their career, the
easier they find each section of a paper to read. An interesting point is how the various career
stages view the importance of each section. Methods, Results and figures seem to be very
important, ostensibly because as academics they have greater skill in their field, allowing them
to be critical of a paper’s methods. It also means they know their field very well, thereofore, the
introduction and abstract have less importance. Early stage PhD students find the methods,
results, and figures fairly difficult to understand. This makes perfect sense as those are the
areas of a paper that require the most knowledge of a field to get through. You are likely to have
a similar experience.
What is it exactly that makes going through this process so difficult and time consuming?
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Authors tend to assume significant background knowledge from readers
Academic syntax is dense and thus difficult for readers to parse
Mathematical expressions are typically condensed and equations reordered for
concision, often skipping steps in derivations
Substantial knowledge gaps are filled if a reader has read cited papers (sort of like —
you need experience to get a job, but need a job to get experience!)
Not all conclusions drawn are correct. Small sample size and power, poor study design,
researcher bias, and selective reporting ensures that you must be a critical reader!
Clearly there is a lot to consider when reading a paper. Scared? Time to lighten the
mood. Here is a hilarious article written on the horrors of reading papers by Dr. Adam
Ruben from Science. It shows even scientists can agree that papers are both difficult to
read and given how dense they are, will keep you regular.
Think about this, the more papers you read, the more you will learn and the faster this process
of reading becomes. Trends start cropping up into plain view, and you begin to gain insight into
the scientific method, understand what certain authors and groups are working on, and form an
appreciation for the field you are learning about. Over time all of this knowledge and skill builds
into your ability to read papers quicker, more efficiently and with greater success. Learning to
read papers is akin to learning to eat. It is messy at first, and your palette is not very well
developed. But over time your eating experience enhances and you learn more about what you
like and don’t like and when a chef’s meal is good and poor.
How Papers are Organized
Good news here. The overwhelming majority of papers follow, more or less, the same
convention of organization:
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●
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Title: Hopefully catchy, possibly sexy! Includes additional info about the authors and their
institutions
Abstract: High level summary
Introduction: Background info on the field and related research leading up to this paper
Methods: Highly detailed section on the study that was conducted, how it was set up,
any instruments used, and finally, the process and workflow
Results: Authors talk about the data that was created or collected, it should read as an
unbiased account of what occurred
Discussions: Here is where authors interpret the results, and convince the readers of
their findings and hypothesis
References: Any other work that was cited in the body of the text will show up here
Appendix: More figures, additional treatments on related math, or extra items of interest
can find their way in an appendix
Developing a Systematic Approach
When you sit down to read it’s important to have a plan. Simply starting to read from page one
to the end will probably do you no good. Beyond retaining limited information, you will be
exhausted and have gained very little for the tremendous effort. This is where many people
stop.
Do plan to spend anywhere from 3–6 hours to really digest a paper, remember they are very
dense! Be ready and willing to make several passes through the paper, each time looking to
extract different information and understanding. And please, do yourself a favor and do not read
the paper front to end on your first pass.
Below are two lists. (i.) the systematic approach I take, more or less, when reading a paper (ii.)
a general list of questions I try to answer as I go through the paper. I typically add more specific
questions depending on the paper.
Let’s get started!
1. Try to find a quiet place for a few hours and grab your favorite beverage (could be
coffee, tea, or anything really). Nowadays I often find myself working in splendid coffee
shops².
2. Start by reading the title and abstract. Aiming to gain a high level overview of the paper.
What are the main goal(s) of the author(s) and the high level results. The abstract
typically provides some clues into the purpose of the paper. Think of the abstract as
advertisement.
3. Spend about 15 minutes skimming the paper. Take a quick look at the figures and note
any keywords to look out for when reading the text. Try to get a sense for the layout of
the paper and where things are located. You will be referencing back and forth between
the different sections and pages later on, it helps knowing where stuff is located. Try not
to spend time taking any notes or highlighting/underlining anything just yet.
4. Turn your attention to the introduction. The more unfamiliar I am with the paper/field, the
longer I spend in the intro. Authors tend to do a good job consolidating background info
and providing copious amounts of references. This section is usually the easiest to read
and it almost feels like you are reading from a textbook. Take notes of other references
and background info you don’t know or want to examine further.
5. This part is extremely critical. Carefully step through each figure and try to get a feeling
for what they are telling you. When I was an undergrad, my neuroscience mentor gave
me some good advice. Paraphrasing: “Figures contain some of the most important
information in a paper. Authors spend a lot of time creating them and deemed the
information they contain to be important enough to communicate to the reader using a
visual. Pay particular attention to them.” You will not understand all the figures very well
the first time you step through them, but you will gain some idea of what the authors
think is most important, and you will also reveal valuable information about what to pay
attention to when you read the other sections.
6. So far you have probably spent about an hour. Take a break. Walk a bit, enjoy a
croissant!
7. Now you are ready to make a first pass through the paper. This time you should start to
take some high level notes. You will come upon words, and ideas that are foreign to you.
Don’t feel like you need to stop at every thing that does not make sense, you can simply
mark it and move on. Aim to spend about an hour and a half. You don’t want to get
bogged down just yet in all the gory details. The goal of the first pass is to get
acquainted with the paper. Like a first date. Your going to learn about the paper, ask
some good questions, maybe make it laugh. But you don’t want to get into every single
little detail. That is rude. Begin again with the abstract, quickly skim through the
introduction, give the methods section a diligent pass. Pay attention to the overall set up,
the methods section includes a ton of detail you don’t need to scrutinize every part at
this point. Finally, read the results and discussion section aiming for some clarity on the
key findings and how these findings were determined. Remember, the authors are trying
to convince you, the reader, of the merit and findings of their work.
8. Saved by the bell. Take a break, do some jumping jacks and get the blood flowing.
Unless you’re in a coffee shop. Then don’t do that.
9. Now that you have a good overview of the paper you are going to get into the nitty gritty
of the figures. Having read the methods, results and discussion section, you should be
able to extract out more gems from the figures. Find those gems. Aim to spend another
30 minutes to an hour on the figures.
10. You should feel confident in taking a second full pass through the paper. This time you
will be reading with a very critical eye. This pass can take a long time an hour or two,
you can also save this for later in the day, or the following day. Pay particular attention to
the areas you marked as being difficult to understand. Leave no word undefined and
make sure you understand each sentence. This pass you are trying to really learn the
paper. Skim through areas you feel confident in (abstract, intros, results). The focus
should be on shoring up what you did not understand previously, and gaining a
command of the methods section and finally being a critical reader of the discussion
section. The discussion section is where you can consider the authors’
reasoning/rational and take what you learned from reading the paper and weigh it
against evidence supplied in the paper. This section should spark some interesting
questions for you to ask your friends, or colleagues. You can even email the authors of
the paper with an insightful question! It might take them a while to get back to you, but
authors do enjoy having dialogue regarding their research and are typically more than
happy to answer a question for a reader.
11. At this point, you should feel confident talking about the paper with colleagues, thinking
critically about the results, and being able to compare the work to other research in the
field (if you have read other papers). To retain and enforce what you have learned, I
suggest you write about the paper. It can simply be a few paragraphs about what you
learned and the significance of the results. You can reference the list of questions you
were answering as you read through the paper.
As mentioned above, here is a general list of questions to help guide you. If you can answer
these you have a solid understanding of the paper, at least to where you can communicate
intelligently about it to others.
1. What previous research and ideas were cited that this paper is building off of? (this info
tends to live in the introduction)
2. Was there reasoning for performing this research, if so what was it? (introduction
section)
3. Clearly list out the objectives of the study
4. Was any equipment/software used? (methods section)
5. What variables were measured during experimentation? (methods)
6. Were any statistical tests used? What were their results? (methods/results section)
7. What are the main findings? (results section)
8. How do these results fit into the context of other research and their ‘field’? (discussion
section)
9. Explain each figure and discuss their significance.
10. Can the results be reproduced and is there any code available?
11. Name the authors, year, and title of the paper!
12. Are any of the authors familiar, do you know their previous work?
13. What key terms and concepts do I not know and need to look up in a dictionary,
textbook, or ask someone?
14. What are your thoughts on the results? Do they seem valid?
I recommend finding people either in person or online to discuss the paper. Start a journal club
with a goal of getting through 1–2 papers a month. The amount of extra insight I have gained
from discussing a paper with a friend is immense. Remember.. the only thing better than
suffering through a paper alone, is suffering through it with friends!
On another note there was a good article written by Keshav³ on how to read a paper. He
introduces and explores a three phrase approach that might be of some interest to you. Give it a
read as well!
Tools to Help You Get the Job Done
You can find papers primarily from several sources:
arXiv: is an open-access repository (maintained at Cornell) where you can freely download and
read pre-print research papers from many quantitative fields. Here is some more general info
about arXiv. Many papers you find on the web will link back to the arXiv paper.
PubMed: They say it best: “PubMed Central® (PMC) is a free full-text archive of biomedical and
life sciences journal literature at the U.S. National Institutes of Health’s National Library of
Medicine (NIH/NLM).” PubMed has a robust search feature if you are looking for medical or life
science related papers.
Google Scholar: I use google scholar just as I would use google. Simply search for a topic,
author or paper and google gets to work, on your behalf. As Google puts it “Google Scholar
provides a simple way to broadly search for scholarly literature. From one place, you can search
across many disciplines and sources: articles, theses, books, abstracts and court opinions, from
academic publishers, professional societies, online repositories, universities and other web
sites. Google Scholar helps you find relevant work across the world of scholarly research.”
Social media: I find out about a lot of new papers simply by following and keeping up with
several people who actively publish. Added bonus.. they typically push other papers they find
interesting and which you might want to know about or read.
Friends and colleagues: Find people interested in the same stuff as you, read papers with them
and learn from each other. I get recommendations for good papers from friends. They act as
good filters.
University: going to your local college or university (if there is one close by) gives you access to
libraries, librarians (very helpful search wizards!) and many journals where you can find and
read articles that are typically behind online paywalls.
As you begin to read more papers you are going to want to store them somewhere. Tossing
PDFs into a folder on your drive is all well and good, but there are creature comforts missing.
Most researchers and grad students use a reference manager. Zotero and Mendeley are very
popular, I like Zotero. Recently, I have been using PaperPile. I like PaperPile because it is
lightweight, lives in my browser, and uses Google Drive to back up and store all my PDFs. It has
a simple, refreshing user interface, and it has a really good tagging and folder hierarchy system.
I can also annotate PDFs in my browser and build citation lists when I write. You get a lot of
these features with almost any reference manager, but I happen to like PaperPile best.
A reference manager will quickly become your best friend as you collect and read more and
more papers.
Thanks for reading through this. I hope you found it helpful and it gave you some good ideas
when tackling your next paper. Most people have their own unique process when reading a
paper. I am sure you will develop your own tweaks in time, hopefully this is a good template for
you to get started.
For now just trust the process.
I am also hoping that we will get some good feedback and comments with other tips and tricks
from readers.
Cheers,
Kyle
Reach me at: datasci@kmshannon.com
linkedin.com/in/kmshannon/
twitter.com/ButFirstData
[1] Hubbard, K. E., & Dunbar, S. D. (2017). Perceptions of scientific research literature and
strategies for reading papers depend on academic career stage. PloS one, 12(12), e0189753.
[2] Shout out to Chris at CoffeeCycle! Simply the best coffee in San Diego.
[3] Keshav, S. (2007). How to read a paper. ACM SIGCOMM Computer Communication Review,
37(3), 83–84.