When media, such as this site wants to explain just how dangerous large animals are, they often quote bite-forces as pounds per square inch (psi).
However, it appears that they mixed up the units, as the numbers they give are the total force in pounds.
But what animals have the strongest bite force per unit area? There is no obvious scaling law here: scale up an animal 2 fold and the cross-sections of both the muscles and the teeth and jaws both quadruple. This puts tiny insect mandibles are on a level playing field with our largest beasts.
Instead, bite-pressure will be niche-dependent as well as the details of the animal's anatomy (such as mechanics of the jaws). It may be somewhat hard to define contact area given the complex shape of teeth and mouthparts in general; maybe one could measure the indented area in a tough material that the animal chomps down on.
Given that our strongest biomaterial found yet is microscopic teeth and that microscopic materials tend to be stronger than bulk materials, it seems likely to be a small animal. Is there any data to support this?
Animals with very high bite pressures - Biology
The laboratory mouse, Mus musculus, belongs to the order Rodentia and family Muridae. The mouse is probably the most genetically and biologically characterized mammal in the world. Mice were first used for the study of respiration in the 17th century. Later in the 19th and 20th century they were bred for their coat color, and subsequently characterized genetically. Most of the common laboratory strains were developed in the early 1900s.
The mouse has short hair, a long naked tail, rounded erect ears, protruding eyes, a pointed snout and five toes on each foot. Mice come in a variety of colors. Insert picture of a mouse, get those cool ones.
Mice have a pair of incisors and three pairs upper and lower of molars. Molars are permanently rooted while the incisors have an open root and grow continuously. Due to this continuous growth of the incisors mice can have problems with overgrown teeth when the upper and lower incisors do not meet properly (malocclusion). Malocclusion can be hereditary or follow trauma, disease or inappropriate diet and/or soft food. There is no permanent cure for overgrown teeth the only treatment is to trim the teeth every 2-3 weeks, if malocclusion persists. Insert pictures.
Mice have a large horseshoe-shaped Harderian gland deep within the orbit. Secretions from the gland contain varying amounts of reddish-brown porphyrin pigment depending on the physiologic state, age, strain and sex of the mouse. The amount of secretions increases during stress and appears as 'red crusts' around the eyes and nostrils.
Normative Values for Mice
Males 20-30 g, Females 18-35g
Mice are communal animals, which live in a very hierarchical society. Within these groups they will aggressively defend their territories. Mums with newborns will likewise aggressively defend their pups and territories. Mice are nocturnal animals but adapt to their environments. While mice are timid, they may bite. Most of mouse behavior is pheromone driven.
Mice are communal animals with a social hierarchical system. This hierarchical system creates a situation whereby the dominant animals often barber (chew off the hair) those of lower rank. This behavior is especially evident among females and is more common in some strains than others. Subordinate animals may have their whiskers, trunk or flank hair removed. There are some suggestions that it may be a cooperative and/or learnt behavior. It appears to be of no sequelae for the health of the mice. Separating the dominant mouse often leads to rise of another dominant mouse. It is important that barbering is differentiated from other causes of hair loss or skin problems e.g. mites, fungi or bacteria by a veterinarian.
Mice have two distinct cervices and uterine bodies. There are separate urethral and vaginal openings. There is a vaginal closure membrane, which is lost at puberty. The inguinal canal remains patent throughout life. Mice have an os penis or os clitoridis associated with external genitalia.
Mice have a 4-5 day estrous cycle, divided into characteristic phases: proestrus, estrus and metestrus. The stage of the estrous cycle can be determined by vaginal cytology. Ovulation occurs at the end of metestrus. Mating leads to formation of a vaginal plug. Plugs persist for 16-24 hours and may last as long as 48 hours. Pregnancy lasts 19-21 days. Females will build a nest prior to parturition if opportunity is provided. Birth usually occurs at night with 10-12 pups being born. Stretching and hindleg extension are usually signs of impeding birth. Babies are born either head or tail first (breech). The female usually eats the placenta. Delivery lasts 1-4 hours, if labor persists call a veterinarian (5-3713). There is a fertile postpartum estrus. Maternal antibody is transferred to the fetus in utero and to the newborn via colostrum.
The young are born incompletely developed (altricius). They are born hairless and their eyes open after 10-12 days. Young are weaned after 21 days at which time they are 10-12 g. Puberty is attained at 4-6 weeks. Breeding onset is 7 weeks and breeding life is 7-9 months. It may be preferable to replace breeders when they are 6 months old.
Mice are generally fed a diet containing low fiber (5%), protein (20%) and fat (5-10%). Feed may be pelleted or powdered. The pelleted feed is supplied as regular, breeder, certified, irradiated or autoclavable. Mice are usually supplied feed free choice and they eat 4-5 g a day (12 g/100 g body weight/day). Water is supplied free choice and they usually drink 3-5 ml a day (1.5 ml/10 g body weight/day). Water may be supplied using a bottle or automatic waterers, and may be further treated by reverse osmosis, ozone, ultraviolet radiation, hyperchlorination or acidification. Picture caging, types of feed, feeders, watering systems
Mouse rooms are usually maintained at 30-70% relative humidity and a temperature of 18-26ºC (64-79ºF) with at least 10 room air changes per hour. The mice are housed in standard shoebox cages with or without filter tops. Filter tops prevent cross contamination of mice limiting the spread of disease and keep facilities clean. Cages with filter tops may have a slightly higher temperature, relative humidity, carbon dioxide and ammonia than the room air. Microisolator tops provide even a higher level of protection than bonnet type filter tops, since they seal better. Static cages as described above are usually changed one to two times a week depending on cage density and housing style. In ventilated cages air is forced into the cage at up to 60 air changes per hour. This keeps the cage dry and reduces build up of ammonia and carbon dioxide. In such situations cages are changed once every 1-2 weeks. Ventilated cages may be kept positive or negative to room air depending on the study being performed.
Mice are usually provided with some kind of bedding in the shoebox cages. Bedding can be paper, wood shaving, wood chips or corncob. In very rare instances mice are housed on wire floors.
Mice should always be clearly identified on cage cards indicating protocol number, strain, sex, age, supplier, investigator and contact person. Procedures performed on the animal should be clearly indicated. Individual mice can be identified using ear punches, ear tags, tattoos, fur dyes, indelible mark on tail or microchips. Picture with standard ear numbering.
Sex is determined using the anogenital distance. Males have a greater (1.5-2 times) anogenital distance than females as well as a larger genital papilla. In neonatal males the testis may be visible through the abdominal wall. Conspicuous bilateral rows of nipples are visible in females at about 9 days of age. Absence of testicles is not a useful criterion for sexing since the testis is retractable into the open inguinal canal throughout life.
Mice should be acclimatized to handling (gentling) to reduce stress. Always talk quietly, move hands slowly and handle them frequently. Mice should be handled at the base of the tail using your fingers or with forceps. Transfer the mice to a firm surface and apply a scruff hold to the loose skin between the ears with your fingers and forefingers while maintaining a grip on the tail. Do not pull the skin too tightly as the mice can choke, too loose a hold will allow the mouse to turn its head and bite. This hold allows you to examine the under belly and perform other procedures. A variety of restraint devices are available to assist in handling mice.
An adult mouse has a circulating blood volume of about 1.5-2.5 ml (6-8% of the body weight), however in older and obese animals this may be lower. Up to 10% of the circulating blood volume may be taken on a single occasion from a normal healthy animal on an adequate plane of nutrition with minimal adverse effect. Always make sure the animal has recovered safely from the procedure and give warm isotonic fluids. This volume may be repeated after 3-4 weeks. For repeat bleeds at shorter intervals, a maximum of 1% of an animal's circulating blood volume can be removed every 24 hours.
Blood can be collected from several sites in the mouse including tail vein, saphenous vein, retro-orbital sinus, brachial vessels, vena cava or cardiac puncture. Always ensure complete hemostasis before returning the mouse to its home cage.
It may be necessary to warm the tail by exposing it briefly to a heat lamp or placing it in a bowl of warm water. The mouse should be restrained in a device for the collection. Blood can be collected from the tail vein (and artery) by making a snip in terminal =3 mm of the tail with a scalpel or sharp scissors. Stroke the tail gently with thumb and finger to enhance flow of blood into the collection vial. Because of the thermoregulatory function of the tail no more than the distal 3 mm should be taken at a time. At the end of the collection apply pressure to the cut end with a gauze bandage and ensure that blood has completely stopped flowing before returning the mouse to the cage. A small nick can also be made at side of the tail 0.5 -2cm from the tail base to collect blood. A fine gauge needle introduced through the skin at a shallow angle can be used to withdraw blood from the tail vein. Apply a tourniquet around the base of the tail to aid in the collection. A butterfly catheter with only about 5 mm of tubing attached to it (rest cut off) may be used instead of a needle and syringe.
Restrain and extend the hind leg applying gentle downward pressure above the knee joint. This stretches the skin making it easier to shave and immobilizes the saphenous vein. Wipe the shaved area with alcohol or sterile lubricate gel and use a 25-gauge needle to puncture the vein (Vein is highlighted by the dark line in the picture below). If done correctly a drop of blood forms immediately at the puncture site and can be collected in a microhematocrit tube. Gentle pressure over the puncture site or relaxation of the restrainer's grip is usually sufficient to stop the blood flow. The scab at the puncture site can be rubbed off at a later date to allow additional blood collection.
The retrorbital sinus is a system of dilated venous channels at the back of the orbit. Blood can be collected form this area in anesthetized mice using a microhematocrit tube. There should be no movement of the head during the procedure. Pressure down with the thumb and forefinger just behind the eye and pull back on the skin to allow the eyeball to protrude. Position a microhematocrit tube along the inner corner of the eye (medial canthus) beside the eyeball. Insert the tube gently but firmly through the conjunctiva towards the back of the eye along the orbit. Rotate the tube gently as you proceed. Blood should flow freely, if, the tube is properly inserted. Tilt the head slightly downward to improve flow. After collecting the blood withdraw the tube and apply pressure on the closed eyelids to stop any bleeding. Remove excess blood with gauze. Complications include damage to the eye and surrounding tissues.
Blood can be collected from the brachial plexus as a terminal procedure in deeply anesthetized mice. Make a cut through the skin at the side of the thorax into the angle of the forelimb (axilla) to expose the axillary vessels. Transect the vessels and allow blood to pool into the pocket created by tenting the skin. Aspirate the mixed venous arterial blood is into an appropriate receptacle.
Vena cava and abdominal aorta
Blood can be obtained from the posterior vena cava or abdominal aorta in a deeply anesthetized mouse following laparotomy. Approach the vessel at a shallow angle using a fine gauge needle attached to a small syringe. This is a terminal procedure.
Up to 1 ml of blood can be obtained from the heart of a deeply anesthetized mouse in a terminal procedure. The most common approach is to lay the mouse on its back and insert a 25 to 30 gauge needle attached to a 1ml syringe just behind the xiphoid cartilage and slightly left of the middle. The needle should be introduced at 10-30 degrees from the horizontal axis of the sternum in order to enter the heart. Alternatively approach the heart laterally immediately behind the elbow at the point of maximum heartbeat.
Administration of substances
Materials to be administered to mice can be given orally e.g. in water or feed or injected systemically through a variety of routes. The average daily consumption of feed and water for an adult 25 g mouse is 3-5 g and 4 ml respectively. The following volumes can be injected into mice safely (based on 25 g mouse): 2-3 ml subcutaneously, 0.05-0.1 ml intramuscularly (0.03 ml per site), 0.50 ml intravenously, 0.1-0.3 ml into the stomach and 2-3 ml intraperitonealy. Intramuscular injections are usually not recommended in mice because of the small muscle mass. A fine gauge needle should be used to make injections in the anterior thigh muscle. It is good practice to use a new needle each time you perform an injection.
Oral gavage is performed using a ball ended feeding needle. Estimate the distance that the needle needs to be inserted into the mouse (usually from the nose to the first rib) and mark it on the needle. Restrain the mouse with the head and body extended as straight as possible to facilitate introduction of the gavage needle. Introduce the needle in the space between the left incisors and molars, and gently direct it caudally toward the right ramus of the mandible. The mouse usually swallows as the feeding tube approaches the pharynx, facilitating entry into the esophagus. If the animal struggles or appears to be in respiratory difficulty withdraw the tube and begin all over again. Once the desired position is attained, inject the material and withdraw the syringe. Monitor the animal after the procedure to ensure that there are no adverse effects.
Subcutaneous injections are usually made into the loose skin over the neck or flank using a fine gauge needle. Insert the needle 5-10 mm through the skin before making the injection. Lack of resistance to the injection is indicative that you are in the right location. Check for leak back especially if a larger volume is injected.
Intraperitoneal injections are usually made in the lower right quadrant of the abdomen. The mouse is restrained with its head tilted lower than the body to avoid injury to internal organs. After swabbing the lower right quadrant with alcohol, a fine gauge needle is introduced slowly through the skin, subcutaneous tissue and abdominal wall. Withdraw the syringe plunger to ensure that you are not in the bladder or intestines. If nothing is withdrawn inject the material slowly. If you accidentally enter the bladder or intestines withdraw and discard the needle and syringe.
Intravenous injections are usually made into the dorsal tail vein. Warm the tail by immersing it in warm water or placing the animal under a heat lamp. The tail vein is easier to see in non-pigmented mice. A fine gauge needle should be used for this procedure.
Signs of pain in the mouse
For a detailed discussion of pain relief in mice refer to module 2. Generally opioids e.g. buprenorphine or non-steroidal anti-inflammatory agents e.g. acetaminophen, ketoprofen, caprofen, ibuprofen are used to relieve pain. Drugs can be administered in water, in jello, as oral drops, by gavage or injected. Drugs administered in water may be broken down in water, or insufficient quantities may be taken due to poor solubility in water or palatability problems.
In general inhalant anesthetics are safer than injectable anesthetics. Halothane and isoflurane are the safest ones to use. Methoxyflurane is no longer available. Use of ether at Johns Hopkins University is subject to restrictions due to safety concerns. Ketamine and xylazine is a common injectable anesthetic combination. Sodium pentobarbital can be used, but it has a narrow safety margin and is associated with a prolonged recovery period. For details on anesthetic techniques refer to the rodent surgery module.
Euthanasia in mice is most often performed by carbon dioxide asphyxiation or overdose of an anesthetic agent. Use of cervical dislocation or decapitation in absence of deep anesthesia must be scientifically justified. All individuals performing euthanasia must be properly trained. Individuals must also ensure that animals are dead before the carcass is disposed. Exsanguination or opening the thoracic cavity will ensure death. AVMA Panel on euthanasia report
Diseases of mice are usually handled as a herd (colony) health problem rather than on an individual animal basis. The goal is to prevent introduction of a disease into a colony rather than to treat animals after disease outbreak. Disease prevention is practiced by institution of a disease surveillance (sentinel) program based on serological and microscopic diagnosis of problems in a representative sample of animals. Due to the widespread movement of animals all over the world with advent of genetic manipulation of animals, the possibility of introducing disease agents in a colony has markedly increased. The expanded use of genetically modified and immunocompromised animals greatly exacerbates the problem. Furthermore the practice of transplanting tumor material into mice provides a portal where these agents can be introduced into animal, especially if the tumors are not screened for adventitious infectious agents. Some important mouse diseases are discussed below to draw attention to the need to adhere to practices recommended by the veterinary staff to avoid these diseases.
Pinworms (Syphacia and Aspicularis) inhabit the intestine (cecum, rectum, colon) and have a direct lifecycle. The eggs are particularly resistant and survive for a long time in the environment. The disease is usually subclinical being marked in weanlings and immunocompromised animals. Symptoms include poor body condition, rough hair coat, reduced growth rate and rectal prolapse. Infection with pinworms has a negative impact on gastrointestinal, growth, behavioral and immunology studies.
Mites affect the skin of mice and up to 100% of the animals may be affected. Affected animals are scruffy, pruritic (itchy), loose hair and have scratch wounds, which can become infected with bacteria. There are changes in the immune responses of affected animals.
Mouse hepatitis virus
This is a viral disease of mice that affects multiple organs. Weanlings are important in maintaining the disease in a colony. Outbreaks result in widespread deaths in neonates and occasionally weanlings, with or without diarrhea. Mouse hepatitis virus causes a wasting disease and high mortality in immunocompromised animals. Usually 100% of the animals are infected. This disease wreaks havoc in a colony, with disruption in research especially in oncology, transplantation, immunology, gastroenterology, metabolism and transgenic technology.
This viral disease is the worst nightmare in a mouse colony. There have been recent outbreaks at several facilities in United States associated with the injection of mouse sera or tissue culture material containing mouse sera into mice. The disease produces massive die off in adult mice and amputation of the limbs (ectromelia) in surviving animals. Pox lesions (mousepox) appear on the skin. There is conjunctivitis, hair loss, as well as swelling of the liver, spleen and lymph nodes. Ectromelia causes high mortality and drastic measures including depopulation are usually taken to eliminate the disease.
This is a fungal disease affecting a wide range of laboratory animals and humans. The organisms are primarily localized in the lungs but may also involve other organs including the eyes, skin etc. It causes a slowly progressive chronic pneumonia with weight loss and eventually death in a large number of immunocompromised animals. The disease has a severe negative impact in research involving immunocompromised animals, pulmonary function and immunology.
1. Kangal – 743 psi
Pictured above, this large, powerful breed is used in Turkey to guard against animal predators. With the strongest bite force of any domesticated dog, they do it well. Dogs in this breed are friendly with their family. They are good with children.
Healthy dogs, the Kangal, are expected to live over 15 years of age. They make excellent guard dogs.
Kangals are alert, territorial, and defensive. They will need social training and “pack leader” training to be your everyday companion dog. If you want this dog, it should be the only animal in the house.
2. Cane Corso – 700 psi
This guard dog originated in Italy. Like most variations of “Mastiff,” the Cane Corso was a war dog.Dogs in this breed have short hair with minimal shedding making them easy to maintain.
They are also hardy and healthy lot. Intelligent and eager, they are easy to train. Most of these pooches are quiet and calm.
Like other guard dogs, this dog usually does not like other dogs and cats. They usually do fine with the kids in their household, but they will need socialization training starting as a puppy to be on the safe side.
3. Dogue de Bordeaux – 556 psi
The Dogue de Bordeaux is the drooling giant from the movie Turner and Hooch. They are affectionate with members of their family. A bit lazy, these canines don’t need a lot of exercises. Unlike some other Mastiffs, this dog is great with kids and other animals – a true gentle giant.
Did I mention the drool? A lot? While friendly with the family, they are suspicious of strangers. They need a secure fence and socialization training. The Dogue can really pack on the pounds, so watch the diet.
4. English Mastiff – 552 psi
Another gentle giant, these pups, is praised by those who own them. Mastiffs are affectionate and do great in families that have kids. They do make excellent guard and watchdogs, but only if trained to be. Energetic and needing daily exercise, this dog is great for the active family or the large yard.
All Mastiffs are independent and strong-willed. They need a dedicated trainer who is willing to put in the time to be a “pack leader.” Like some other breeds, the English Mastiff can become destructive when left alone too long.
5. Dogo Canario – 540 psi
Also known as the Perro de Presa Canario, this is one ancient war dog still known to be aggressive. Brave and loyal, dogs in this breed make great guard dogs. Great for the active family, these pooches need exercise, a large yard, and mental stimulation. Pups in this breed are said to be friendly if raised with kids and other animals.
This breed needs an experienced trainer. To have a successful relationship with strong dogs like this one, the owner needs to have the time and experience to socialize and train this animal properly.
6. Dogo Argentino – 500 psi
Breed to be a big game hunter, this breed has great reflexes and strong stature. These dogs are clean and shed very little, making them easy to maintain. Active and friendly, these canines are great for a large family. They need time with their family to be the best they can be.
A strong and independent dog, this fur-baby needs a strong, experienced trainer. Socialization and obedience training must start at a young age and stay consistent. This is another breed that must have pack leader training.
Introduction: the occurrence of nociception and pain
The ability to detect dangerous, damaging stimuli is adaptive in terms of survival, and thus the evolution of an early warning system in animals seems intuitive. Indeed, the nociceptive system, which detects noxious, harmful, injury-causing stimuli such as extremes of temperature, high mechanical pressure and irritant chemicals, has been identified in invertebrates (e.g. Drosophila and Caenorhabditis elegans Tobin and Bargmann, 2004 Neely et al., 2010 Im and Galko, 2012) through to humans (reviewed in Sneddon et al., 2014). Nociception is the simple perception of a noxious event and is typically accompanied by a reflex withdrawal response away from the source of damage. In humans, negative ‘feelings’ of discomfort or suffering are experienced alongside the injury and this is termed pain. The concept of pain occurring in animals has been extensively debated, with some authors suggesting only primates and humans can experience the adverse affective component as they possess a human (or similar in primates) neocortex (Rose, 2002 Rose et al., 2014). Opposing this opinion, scientists suggest that the negative experience that accompanies tissue damage is crucial in altering an animal's subsequent behaviour to perform protective and guarding reactions, enabling the animal to avoid such stimuli in future, and for avoidance learning to occur (Sneddon et al., 2014). This implies that the unpleasant internal state of experiencing pain goes hand in hand with its perception as it has to be such a strong aversive stimulus to ensure animals will alter future behaviour and learn from the event. If this were not the case, animals would continue to damage themselves repeatedly, resulting in disease, loss of limbs and even mortality. It is unlikely that animals living in very different environments will have developed the same nociceptive or pain-detecting neural machinery as humans. Evolution and life history place very diverse pressures on different animal groups as well as exposing them to differing types of nociceptive stimuli. For example, fish living in an aquatic world can maintain buoyancy, so the risk of collision due to gravity is likely to be rare compared with a terrestrial vertebrate (Sneddon, 2004). Therefore, evolution, ecology and life history may have shaped nociceptive and pain systems in aquatic animals to meet the demands of their environment in quite a dissimilar way to terrestrial animals (Broom, 2001 Rutherford, 2002).
Studies on the bird brain have challenged old dogma on brain evolution and have shown that the theory of linear and progressive evolution, where the cerebral cortex was believed to have evolved from lower to higher vertebrates, with birds at an intermediate stage, are incorrect (see Jarvis et al., 2005 for review). The avian cerebrum was proposed to be relatively simple in structure and controlled primitive behaviours (Ariëns Kappers et al., 1936). Modern experimentation demonstrates that the avian cortex developed from pallial (or cortical-like), striatal and pallidal regions from a common avian/reptilian ancestor. The avian pallial region is differentially organized compared with the mammalian pallium or cortex in that the avian region is nuclear and the mammalian cortex is layered. However, the pallia of the two groups perform similar functions and have comparable connectivity. Thus, the avian cortical regions are different from the mammalian or human cortex but have evolved analogous roles. Thus, we should expect that other non-mammalian vertebrate taxa may have brain structures that differ from the mammalian cortex anatomically, but that may have evolved to perform similar functions. This example demonstrates how the study of disparate animal groups can enlighten us about the evolution of the nervous system. In this review, the evidence for pain and nociception in wholly aquatic animals will be considered by discussing the differences with terrestrial groups to ascertain how the watery environment may have resulted in anatomical, neurobiological and functional differences. Unfortunately, there are negligible data on aquatic mammals and aquatic birds, and very few data published on aquatic forms of amphibians (see Guenette et al., 2013) therefore, this review will focus upon the aquatic animal groups where there is significant empirical evidence, specifically fish, cephalopods and crustaceans.
Definitions of animal pain have proposed that there are two key criteria animals must fulfil to be considered capable of experiencing pain (Table 1 Sneddon et al., 2014). The first is that whole-animal responses to a noxious, potentially painful event differ from those to innocuous stimulation. More specifically, it is suggested that: animals must have the neural apparatus to perceive damaging stimuli with nociceptors, often free nerve endings that specifically detect harmful stimuli the information is conveyed to the central nervous system (CNS) from the periphery central processing occurs involving brain areas that innervate motivational and emotional behaviour and learning physiological responses are altered that may be linked to a stress response behavioural alterations are not simple reflexes, with long-term responses including protective behaviours and avoidance as part of the response and finally all of these reactions should be reduced by the use of analgesics or painkillers (although note that these may only be effective in mammals and specific compounds may not necessarily work in invertebrates with different receptors, etc. e.g. Barr and Elwood, 2011). The first invertebrate species in which nociceptors were identified was the leech (Hirudo medicinalis), an aquatic annelid inhabiting freshwater (Nicholls and Baylor, 1968). These nociceptors have similar properties to mammalian nociceptors including responding to multiple types of noxious, damaging stimuli, classifying them as polymodal (Pastor et al., 1996). Further studies have demonstrated that analgesic compounds and stimulation of adjacent touch receptors can reduce nociceptor activity (Higgins et al., 2013 Yuan and Burrell, 2013), the latter being comparable with gate control in mammals whereby touch modulates pain transmission (Melzack and Wall, 1965). Therefore, the physiological and molecular mechanisms may be highly conserved between aquatic invertebrates, vertebrates and terrestrial vertebrate groups (Sneddon et al., 2014).
The two key principles and detailed criteria for pain in animals
Blood primarily moves through the body by the rhythmic movement of smooth muscle in the vessel wall and by the action of the skeletal muscle as the body moves. Blood is prevented from flowing backward in the veins by one-way valves. Blood flow through the capillary beds is controlled by precapillary sphincters to increase and decrease flow depending on the body&rsquos needs and is directed by nerve and hormone signals. Lymph vessels take fluid that has leaked out of the blood to the lymph nodes where it is cleaned before returning to the heart. During systole, blood enters the arteries, and the artery walls stretch to accommodate the extra blood. During diastole, the artery walls return to normal. The blood pressure of the systole phase and the diastole phase gives the two pressure readings for blood pressure.
Modeling the bite force of T. rex helps to explain important dietary clues about the animal.
Tyrannosaurus rex needs no introduction. It is the go-to dinosaur for all ages that has been popular since its description in 1905. Partly due to its popularity, scientists have researched and debated much of its life history. One of the more popular debates is centered around what T. rex ate and whether it was a carnivore or a scavenger. While there are many different ideas surrounding what the dinosaur may or may not have eaten, one comparison had not been drawn between Tyrannosaurus and some modern animals – were bones a part of its diet on purpose? In order to answer this question, its jaw and skull anatomy had to be biomechanically modeled to observe how well its musculature, bite force, and tooth structure would have withstood biting down on bone.
Modern reptiles will swallow bones whole rather than break them crushing bones for food is generally thought of as a mammalian trait since most modern reptiles do not have the jaw strength to crush larger bones. However, T. rex was likely able to crush the bones of its prey. To understand this difference, a recent paper revisited how hard Tyrannosaurus rex could bite and how that force influenced its dietary habits. Using jaw muscle modeling, bite force estimation, and tooth resistance to pressures, the researchers attempted to understand how it was possible that T. rex was able to finely crush bones as it ate.
According to the authors of the paper, large-bodied tyrannosaurids were able to crush bone thanks to their enormous bite forces which, in the case of Tyrannosaurus , were calculated to be between 8,526 to 34,522 newtons of force based on its skull and jaw dimensions. To put this into perspective, some crocodilians can bite down with approximately 16,500 newtons of force and we humans can only bite down with a measly 890 newtons. Similarly, the teeth were able to withstand the stresses of repeatedly biting hard bone at high pressures. Most of the maxillary teeth could withstand stresses that fracture the dense outer layer of bones for up to 25 mm from the tip of the tooth. Once the tooth had punctured a bone, the bite force would have easily crushed the remaining part of the bone.
Osteophagy, the tendency of an animal to knowingly ingest bones as part of its normal diet, is one method animals use to gain the nutrients they need and is fairly common throughout all animals. Modern reptiles and birds that break bones for consumption use a variety of methods to access the nutrition locked away in the bones themselves. For example, alligators will use their blunt teeth to bite and puncture other animals (bones included) and the bearded vulture will drop bones from great heights to break them and access the marrow inside. Both birds and reptiles that utilize osteophagy regularly have strong stomach acids with a very low pH – thus allowing for the bones they eat to be easily chemically dissolved. Similarly, tyrannosaurids such as Tyrannosaurus would have been able to break down bone and easily digest them to allow extra nutritional intake. In the case of T. rex, the teeth and jaws were perfect to crack and fragment bones and, based on bone fragments in collected coprolites (fossilized poop that has been associated with T. rex ), easily digest them as well.
Scientists have known for a long time that the bite force of Tyrannosaurus was immense and played a part in its ability to eat. However, the extent of T. rex ’s bite force as a function of diet had not been explored. The combination of bite force, the teeth’s ability to withstand massive forces, and repeated biting in the same area along a bone gave Tyrannosaurus the ability to fragment bone with relative ease. Adaptations to freeing more nutrients from bones would have helped sustain its enormous body size and helped it to be as evolutionarily successful as it was.
This research is dedicated to the late T. Hunter and J. B. Thorbjarnarson. Both provided invaluable insights and assistance for our study. A special thanks goes out to D. C. Drysdale for generously allowing scientific access to the entirety of his crocodilian specimens at the St. Augustine Alligator Farm Zoological Park. We also thank the curatorial staff members of St. Augustine Alligator Farm Zoological Park and Crocodylus Park. J. Gatesy generously allowed access to his comprehensive data on the phylogeny for the Crocodylia. C. Brochu and Wann Langston provided helpful discussions about fossil crocodilian specimens, historical size range, systematics, and taxonomy. S. Deban assisted with the gathering and analysis of data on underwater jaw closure. K. Womble produced the graphics.
T-Rex's bone-shattering bite was much worse than its bark
Few animals capture the imagination quite like the Tyrannosaurus rex – even if it wasn't the biggest predator to ever stalk the Earth, and it may have sounded more bird than beast. But a new study shows that the King may still deserve its crown, thanks to a terrifying set of jaws that could deliver one of the strongest bites of any land animal in history. It's an ability that T-rex probably used to pulverize and eat the bones of its unfortunate prey.
A group of scientists from Florida State University began by studying the bite force of crocodiles and alligators, which are at the top of today's list of animals you don't want to find yourself between the jaws of. In 2012, the team found that a saltwater crocodile packed a bite of 3,700 pound-force (16,460 newtons), By comparison, a human vigorously tearing into a steak tops out at around 200 lbf (890 N).
Using the jaw musculature of modern crocodiles, Florida State University researchers calculated that the T-rex might have had a bite force of 8,000 lb (35,586 N), and 431,000 psi
Using the jaw musculature of crocodiles as a starting point, the Florida State researchers compared the reptilian figures with those of birds, a closer modern relative of dinosaurs, to create a model for the biting power of a T-rex. The dinosaur, they found, could chow down with about 8,000 lbf (35,586 N), more than twice the force of a saltwater croc. But the team points out that in practical terms, there's more to the story than that one number.
"Having high bite force doesn't necessarily mean an animal can puncture hide or pulverize bone, tooth pressure is the biomechanically more relevant parameter," says Gregory Erickson, co-author of the study. "It is like assuming a 600 horsepower engine guarantees speed. In a Ferrari, sure, but not for a dump truck."
Tooth pressure is determined by how the shape of the teeth focuses the pressure into a smaller point, and the T-rex's long, cone-shaped chompers were perfect for piercing flesh and shattering bone. Channeling the force, a Tyrannosaurus bite could impart as much as 431,000 pounds per square inch (psi), which helped the animal pulverize the bones of its prey to give it a nutritional advantage over other predators of its day. To compare, saltwater crocs were measured at 360,000 psi.
"It was this bone-crunching acumen that helped T-rex to more fully exploit the carcasses of large horned-dinosaurs and duck-billed hadrosaurids whose bones, rich in mineral salts and marrow, were unavailable to smaller, less equipped carnivorous dinosaurs," says Paul Gignac, co-author of the study.
This section of Triceratops pelvis shows some 80 T-rex bite marks, indicating a "puncture and pull" pattern of biting
This feeding style is very different from modern crocodiles, which bite to kill before swallowing their meals mostly whole. Due to their tooth structure, as well as bite marks found on Triceratops bones, the researchers suggest that the T-rex ate with a biting and chewing pattern, in the style of modern mammals like hyenas and wolves. That means this feeding pattern arose earlier than previously thought.
The Tyrannosaurus Rex’s Dangerous and Deadly Bite
Tyrannosaurus rex has always been recognized as fearsome—the New York Times labeled it the “prize fighter of antiquity” when the first mounted T. rex bones were displayed in 1906—but thanks to two British researchers, it’s now clear that the giant carnivore bit harder than experts had thought. A lot harder.
From This Story
The Tyrannosaurus Rex known as Stan, excavated in South Dakota in 1992, is one of the most complete tyrannosaurus rex skeletons in the world. (Greg Latza / AP Images)
Karl Bates, a biomechanics expert at the University of Liverpool, and Peter Falkingham, a paleontologist at the Royal Veterinary College, London, and Brown University, acknowledge that measuring the biomechanics of an extinct species “is notoriously difficult and involves numerous assumptions.” But for their assessment of T. rex’s bite, published in Biology Letters, they constructed a three-dimensional digital model of the animal’s skull and reconstructed the relevant jaw musculature, based on anatomical research on birds (which are, after all, living dinosaurs) and crocodilians (the closest living cousins to Dinosauria as a group). Previous assessments relied on extrapolations from crocodile bites or fossil T. rex tooth marks.
When Bates and Falkingham used computer models to simulate T. rex’s bite, the result was “quite surprising,” Bates told us: a maximum bite force of almost 12,800 pounds, about the equivalent of an adult T. rex’s body weight (or 13 Steinway Model D concert grand pianos) slamming down on its prey. That would make T. rex the hardest-biting terrestrial animal ever known. (C. megalodon, an extinct giant shark, bit at an estimated 41,000 pounds Deinosuchus, an ancient crocodilian, at about 23,000 pounds.) Bates and Falkingham’s figure was two to three times greater than previous estimates, six to seven times greater than the biting force they calculated for the dinosaur Allosaurus and about three and a half times greater than the hardest bite measured in an extant species, an Australian saltwater crocodile.
“The posterior part of the skull that housed the muscles was particularly large,” Bates says. Rare juvenile T. rex skeletons indicate that these dinosaurs were leggy runners with relatively shallow skulls incapable of anchoring muscles that would generate a bite proportional to the adults’. In Bates and Falkingham’s tests, juvenile T. rex bites topped out at about 880 pounds. But as the animal matured, its jaw-closing muscles grew exponentially, to the point where they were huge “even for an animal of its colossal size.”
Young T. rex were still formidable—they just targeted different prey. While the juveniles raced down small game, Bates notes, the adults had the power to bring down megaherbivores such as Edmontosaurus and Triceratops. This phenomenon, called resource partitioning, would have reduced competition between parents and offspring—a big evolutionary advantage. As if T. rex needed it.
About Riley Black
Riley Black is a freelance science writer specializing in evolution, paleontology and natural history who blogs regularly for Scientific American.
T. Rex's Bite More Dangerous Than Previously Believed
The tyrant lizard, also known as Tyrannosaurus rex, had the strongest bite of any known land animal, new research suggests.
"Our results show that the T. rex had an extremely powerful bite, making it one of the most dangerous predators to have roamed our planet," study researcher Karl Bates, of the University of Liverpool, said in a statement.
Younger T. rexes didn't have such strong bites, the researchers found, which probably meant they had a different diet and relied less on the fearsome bite than their older counterparts. This differing diets likely led reduced competition between generations of T. rex, the researchers said. [Image Gallery: The Life of T. Rex]
The new estimate of bite force is higher than past estimates that relied on indent measures in which they pressed down the skull and teeth onto a bone until they got the imprints that matched those on fossils. In the new study, the researchers created a computer model of the dinosaur's jaw by first digitally scanning skulls from an adult and juvenile T. rex, an allosaurus, an alligator and an adult human. They used these scans to model each animal's bite.
"We took what we knew about T. rex from its skeleton and built a computer model," Bates said. "We then asked the computer model to produce a bite so that we could measure the speed and force of it directly."
The force exerted at one of T. rex's back teeth would have been between 7,868 and 12,814 pounds-force (35,000 and 57,000 newtons). This force would be akin to having a medium-size elephant sit on you.
Young vs. old
The shape of T. rex's skull allowed room for lots of muscles, creating what is "by far the highest bite forces estimated for any terrestrial animal," the researchers write in the paper, to be published tomorrow (Feb. 29) in the journal Biology Letters, but it is possible the extinct gigantic shark Megalodon had a stronger bite.
"If you consider that the lion and alligator [bite strength] are so much lower (as reported in our paper), and think of what they can bite through, that can give you a sense of the power in a T. rex bite," study researcher Peter Falkingham, of the University of Manchester in the United Kingdom, told LiveScience in an email. "Such a powerful bite may have enabled T.rex to crush large bones."
(Past research has suggested T. rex's fused nasal bones boosted its bite force, while also keeping the predator's skull from breaking from a serious chomp.)
Even when Falkingham and colleagues scaled the models for body size differences, this bite was relatively much stronger than the bite of a juvenile T. rex. In its early years of life, T. rex's bite was weaker, but the young dinosaurs might have also been more athletic and had longer arms in proportion to their body size, previous research has suggested.
These differences could mean that the dinosaur's diet would have changed over time &mdash starting on smaller prey, but growing into a ferocious predator to even the largest animals as it matured. These dietary differences would have reduced competition between older and younger T. rexes, Falkingham said.
You can follow LiveScience staff writer Jennifer Welsh on Twitter @microbelover. Follow LiveScience for the latest in science news and discoveries on Twitter @livescience and on Facebook.