Why is the human body able to repair a broken bone and not a heart muscle?

Why is the human body able to repair a broken bone and not a heart muscle?

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The human body can repair skin/organ laceration, fractures, even repair nerves - albeit the duration and rate of recovery differ.

Why can the heart muscles not repair themselves?

Part of the answer may be(*) that in bone, you have still stem cells everywhere (blood capillaries that grow until checked and osteoblasts), i.e., also in the vicinity of the wound. The same applies to skin, by the way, where the epidermis continues to produce skin layers until eternity. This facilitates repair enormously, as new cells won't have to move far to take their place.

Contrast this with heart or spinal cord, where the placement of the cells took place in early development when distances were small and most cells were undifferentiated. Also, no tissue like epidermis was left in the vicinity that could act as a near source of new cells.

(*) I said "may be" because 1. I have no ref for this, and 2. I have no idea to what extent other factors are involved. I would be thankful for counterexamples.

The heart does have stem cells in it, and there is cell turnover in the heart, of about 1% per year. Which is much slower than your skin, but not nothing. This allows your heart to grow during your life, and remodel itself slightly to become stronger/more efficient when you get in shape.

The heart can repair itself, when damaged it doesn't simply stay damaged. Unfortunately, the 'repair' leaves particularly useless scar tissue. After a heart attack, the dead muscle does repair itself, but very poorly. This 'scar' barely contracts, and isn't as strong as the heart wall around it.

This is mostly a function of the very specialized heart myocytes, and the evolutionary (relative) uselessness of being able to regenerate your heart after injury. In the wild, if your heart was injured, you were probably dead.

Part of the answer is in fact extremely simple: coronary arteries (irrigating the heart muscle) are terminal vessels. This means that when a coronary artery sustains damage, the area it was irrigating becomes biologically dead with a very limited potential for recovery. The affected area turns to fibrous tissue produced by cells with much lower oxygen requirements than cardiomyocytes.

A fractured bone is not dead at all, in most cases. Put a fracture under a microscope, and you will see plenty of cells performing heavy duty. That is because the area, although damaged, still receives nutrients and at least some blood.

However, there are special cases:

  • bone that is terminally vascularized: such bone regions will die when fractured, since they receive no more nutrients and oxygen. The classic example is fracture of the scaphoid bone in the wrist. Some types of fracture in this bone shear the nourishing artery, and the bone dies off instead of healing.

  • slowly progressive ischemia due to atherosclerosis: when a coronary artery progressively gets clogged instead of this being a sudden event, the cells inside the danger territory secrete messengers that will activate new vessel growth. This phenomenon is called collateralization, and will allow a subset of the cells at risk to survive when the coronary artery gets completely clogged.

Now about the stem cell issue: I seem to remember that stem cells do exist in the heart, but this is less pertinent in this case, since even stem cells need oxygen and nutrients to survive.

Ignoring the actual physiological mechanism(s) for why, consider that from a natural selection point of view, there probably isn't as much incentive for a body to heal itself of a condition that largely affects post-reproduction age individuals. While healing a broken bone would likely allow a young person to further reproduce, healing a heart ailment would have a disproportionately lower chance of doing so.

In simple terms, heart injury, most likely to be myocardial infarction (heart attack) involves death of the vascular heart tissue. Thus the word infarction. Death of a tissue just simply leads to the production of a fibrous scar. Therefore the heart muscle cannot repair that area of injury to 100% as before.

How do broken bones heal?

If you experience engine trouble, you take your car to a mechanic. If your pipes leak, you call a plumber. And if you fracture a leg, the usual course of action is to visit a doctor. But unlike other things that may break in life, bones begin healing on their own before you even set foot in a waiting room.

The human body possesses amazing healing powers that enable it to bounce back from a vast array of illnesses and injuries. Sometimes broken bones can heal so thoroughly within a few months that even an x-ray can't determine the original fracture line.

Doctors often play a vital, sometimes lifesaving, role in a bone's healing process. But, these experts basically help the body heal itself. Doctors provide optimal conditions for bone repair and healing to take place. The rest is up to your cells.

But how does this amazing biological process work? How can a fractured limb grow back to its former strength? To understand, you first have to take a closer look at just what bones are made of and how alive they really are.

How Broken Bones Repair Themselves

It's easy to think of our bones as solid, lifeless matter where all of our living tissue just sits. But your skeleton is as much a living part of your being as your softer tissues and organs. The body stores minerals in the hard, compact bone. It produces red blood cells in the inner red marrow and stores fat in the yellow marrow.

It's important to remember that your bones are constantly changing. Cells called osteoclasts constantly break down old bone so that osteoblasts can replace it with new bone tissue -- a process called bone remodeling. Another type of cell called a chondroblast forms new cartilage. These are three of the primary cells responsible for bone growth -- and not just the bone growth you experience early in life. This constant bone remodeling gradually replaces old bone tissue with new tissue during the course of months.

Almost immediately after the break, the body begins to try and put itself back together again. Doctors often divide the overall process into four phases:

  1. When a bone breaks, the fissure also severs the blood vessels running down the length of the bone. Blood leaks out of these veins and quickly forms a clot called a fracture hematoma. This helps to stabilize the bone and keep both pieces lined up for mending. The clot also cuts off the flow of blood to the jagged bone edges. Without fresh blood, these bone cells quickly die. Swelling and inflammation follow due to the work of cells removing dead and damaged tissue. Tiny blood vessels grow into the fracture hematoma to fuel the healing process.
  2. After several days, the fracture hematoma develops tougher tissue, transforming it into a soft callus. Cells called fibroblasts begin producing fibers of collagen, the major protein in bone and connective tissue. Chondroblasts then begin to produce a type of cartilage called fibrocartilage. This transforms the callus into a tougher fibrocartilaginous callus, which bridges the gap between the two pieces of bone. This callus generally lasts for about three weeks.
  3. Next, osteoblasts move in and produce bone cells, transforming the callus into a bone callus. This hard shell lasts three to four months, and it provides necessary protection and stability for the bone to enter the final stage of healing.
  4. At this point, the body establishes the position of the bone within the flesh, begins reabsorbing bits of dead bone, and creates a hard callus to bridge the gap between the two pieces of bone. However, this bulge of tissue needs a lot of work before the bone can take any strain. Osteoclasts and osteoblasts spend months remodeling bone by replacing the bone callus with harder compact bone. These cells also decrease the callus bulge, gradually returning the bone to its original shape. The bone's blood circulation improves and the influx of bone-strengthening nutrients, such as calcium and phosphorus, strengthen the bone.

But even in the best of cases, fractures often require medical attention to heal as smoothly as possible. Did Humpty Dumpty just need the help of a talented orthopedist? Learn how doctors help broken bones heal properly on the next page.

Are you at risk for a broken bone? The American Academy of Orthopaedic Surgeons reports that men under the age of 45 and menopausalwomen over the age of 45 experience the highest number of breaks. Your wrist's radius bone is the most commonly broken bone, at least if you're younger than the age of 75. Among the elderly, hip fractures are far more common [source: OSF Healthcare].

Types of Fractures

Fractures are classified by their complexity, location, and other features (Figure 6.5.1). Table 6.4 outlines common types of fractures. Some fractures may be described using more than one term because it may have the features of more than one type (e.g., an open transverse fracture).

Figure 6.5.1 – Types of Fractures: Compare healthy bone with different types of fractures: (a) open fracture, (b) closed fracture, (c) oblique fracture, (d) comminuted fracture, (e) spiral fracture , (f) impacted fracture, (g) greenstick fracture, and (h) transverse fracture.

Types of Fractures (Table 6.4)
Type of fracture Description
Transverse Occurs straight across the long axis of the bone
Oblique Occurs at an angle that is not 90 degrees
Spiral Bone segments are pulled apart as a result of a twisting motion
Comminuted Several breaks result in many small pieces between two large segments
Impacted One fragment is driven into the other, usually as a result of compression
Greenstick A partial fracture in which only one side of the bone is broken, often occurs in the young
Type of Fracture Description
Open (or compound) A fracture in which at least one end of the broken bone tears through the skin carries a high risk of infection
Closed (or simple) A fracture in which the skin remains intact

The Story of Biomaterials

Biomaterials have improved significantly since they were first developed, and they are still changing, as scientists continue to understand more about diseases and how the biomaterials interact with the body [3]. Biomaterials can be created from a variety of materials, depending on what they will be used for [2, 3]. For example, they can be made from various types of natural components, such as collagen which is found in the body or alginate which comes from seaweed, synthetic materials, such as metal, or a combination [2, 3]. The earliest biomaterials did not interact with the human body, but they had similar physical properties to the damaged organs they were being used to repair or replace. These materials were often created from various metals, ceramics, or substances, such as rubber. These early biomaterials were commonly used as prosthetics, which are artificial body parts such as a leg or a heart, but they had poor combability with the body so they would often be rejected. These early materials could not interact with the body on a cellular level, which is what today’s biomaterials aim to do. Advances in the development of novel biomaterials have led materials that can interact with the body to promote healing and regeneration . These newer biomaterials were considered bioactive, meaning they could interact with the body, and form chemical bonds with tissues. This is seen in hip implants that promote bone growth which allow a calcium layer, called hydroxyapatite, to grow on the implant. The newest biomaterials, known as third generation biomaterials, are made to interact with the body and cause a specific response from cells in the body. Third generation biomaterials can also mimic the body’s natural 3D structure and stimulate tissue regeneration (re-growth) [3]. Scientists have improved biomaterials tremendously and they continue to work on them, changing the materials’ properties to operate more effectively in patients’ bodies.

6.5 Fractures: Bone Repair

A fracture is a broken bone. It will heal whether or not a physician resets it in its anatomical position. If the bone is not reset correctly, the healing process will keep the bone in its deformed position.

When a broken bone is manipulated and set into its natural position without surgery, the procedure is called a closed reduction . Open reduction requires surgery to expose the fracture and reset the bone. While some fractures can be minor, others are quite severe and result in grave complications. For example, a fractured diaphysis of the femur has the potential to release fat globules into the bloodstream. These can become lodged in the capillary beds of the lungs, leading to respiratory distress and if not treated quickly, death.

Types of Fractures

Fractures are classified by their complexity, location, and other features (Figure 6.20). Table 6.4 outlines common types of fractures. Some fractures may be described using more than one term because it may have the features of more than one type (e.g., an open transverse fracture).

Type of fracture Description
Transverse Occurs straight across the long axis of the bone
Oblique Occurs at an angle that is not 90 degrees
Spiral Bone segments are pulled apart as a result of a twisting motion
Comminuted Several breaks result in many small pieces between two large segments
Impacted One fragment is driven into the other, usually as a result of compression
Greenstick A partial fracture in which only one side of the bone is broken
Open (or compound) A fracture in which at least one end of the broken bone tears through the skin carries a high risk of infection
Closed (or simple) A fracture in which the skin remains intact

Bone Repair

When a bone breaks, blood flows from any vessel torn by the fracture. These vessels could be in the periosteum, osteons, and/or medullary cavity. The blood begins to clot, and about six to eight hours after the fracture, the clotting blood has formed a fracture hematoma (Figure 6.21a). The disruption of blood flow to the bone results in the death of bone cells around the fracture.

Within about 48 hours after the fracture, chondrocytes from the endosteum have created an internal callus (plural = calli) by secreting a fibrocartilaginous matrix between the two ends of the broken bone, while the periosteal chondrocytes and osteoblasts create an external callus of hyaline cartilage and bone, respectively, around the outside of the break (Figure 6.21b). This stabilizes the fracture.

Over the next several weeks, osteoclasts resorb the dead bone osteogenic cells become active, divide, and differentiate into osteoblasts. The cartilage in the calli is replaced by trabecular bone via endochondral ossification (Figure 6.21c).

Eventually, the internal and external calli unite, compact bone replaces spongy bone at the outer margins of the fracture, and healing is complete. A slight swelling may remain on the outer surface of the bone, but quite often, that region undergoes remodeling (Figure 6.21d), and no external evidence of the fracture remains.

Interactive Link

Visit this website to review different types of fractures and then take a short self-assessment quiz.

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    We’re creating fully functional tissue which already beats and contracts – Sanjay Sinha

    “We believe that these patches will stand a much greater chance of being naturally assimilated into a patient’s heart, as we’re creating fully functional tissue which already beats and contracts through combining all these different cell types which communicate with each other,” Sinha says.

    “We know the epicardium cells are particularly important in co-ordinating proper development of heart muscle because research has shown that in developing embryos, there’s a lot of crosstalk which occurs between the epicardium and the developing heart.”

    Watch a heart patch beat in this video from Stanford University below:

    Bones Facts for Kids

    • At birth, you have 270 bones in your body.
    • Adults have 206 bones as they fuse together as you get older.
    • You have 26 bones in your foot.
    • You have 27 bones in your hand.
    • Some bones help protect important organs in the body, for example, the rib cage helps to protect your heart, liver and lungs.
    • Our body has 5 different types of bones flat, short, long, irregular and sesamoid.
    • The inside of her bones is filled with marrow which is a soft tissue.
    • When bones are connected they form a joint. There are several different types of joints ball and socket, pivot, hinge and saddle joints.
    • The brain is protected by 8 different types of bones.
    • The longest bone in the body is the ‘femur’. This is the thigh bone and is also considered to be the strongest bone in the body. report this ad
    • The smallest bone in the body is actually located in your middle ear. It is called the ‘staples’ (or stirrup) and it’s only 0.11 inches (2.8 mm) long.
    • Like many parts of the body, our bones are constantly being worked and worn down and then re-made again. It takes seven years for the cells in the bones to regenerate so literally every 7 years we have a new bone.
    • There are 54 bones in your hands, fingers and wrists. This is the part of the body that has the most bones.
    • Although our teeth are part of the complete skeleton, they are not considered bones.

    Male and female skeletons have a few differences. The male skeleton is typically larger and the female skeleton typically smaller. Female skeletons also have a different shape to the pelvic bone so that it is wider and angled to allow childbirth.

    Almost all of our bones have a strong outer layer that is dense with an inside layer that is spongy and partially filled with air. The very center is flexible, soft tissue that is called the ‘bone marrow’.

    4% of the human body mass is made up of bone marrow. It is the marrow that produces the red blood cells that we need to carry oxygen throughout our body. Bone marrow also creates important components of our lymphatic system in the production of lymphocytes. These help with our immune system.

    Both calcium and vitamin D are important to help in keeping healthy, strong bones.

    The parts of the skeleton where our bones meet are called ‘joints’. The joins allow movement, with the exception of the cranium joints. The bones are held in place at our joints with ligaments and muscles. Cartilage is another type of tissue that covers the surface of each bone join to keep the bones from rubbing against each other.

    Hands-on Activity Repairing Broken Bones

    Units serve as guides to a particular content or subject area. Nested under units are lessons (in purple) and hands-on activities (in blue).

    Note that not all lessons and activities will exist under a unit, and instead may exist as "standalone" curriculum.

    • Biomedical Engineering and the Human Body
      • Engineering Bones
        • Prosthetic Party: Build and Test Replacement Legs
        • Sticks and Stones Will Break That Bone!
        • Muscles, Oh My!
          • The Artificial Bicep
          • Measuring Our Muscles
          • Body Circulation
            • Clearing a Path to the Heart
            • Breathe In, Breathe Out
              • Polluted Air = Polluted Lungs
              • Digestion Simulation
                • Protect That Pill
                • My Mechanical Ear Can Hear!
                  • Sounds All Around
                  • Biomedical Devices for the Eyes
                    • Protect Those Eyes
                    • We've Come a Long Way, Baby!
                      • You're the Expert
                      • DNA: The Human Body Recipe
                        • DNA Profiling & CODIS: Who Robbed the Bank?
                        • DNA Build
                        • Bone Fractures and Engineering
                          • Repairing Broken Bones
                          • Living with Your Liver

                          TE Newsletter

                          Devices such as plates, screws and rods are designed by engineers for doctors to surgically implant to help mend severely fractured bones.


                          Engineering Connection

                          Biomedical and materials engineers create devices that doctors use to repair severe bone fractures. Materials engineers develop biocompatible materials that integrated into the body easily. Biomedical engineers use these materials to design pins, plates, rods and screws that can be used to help support and repair broken bones.

                          Learning Objectives

                          After this activity, students should be able to:

                          • Describe how engineers aid doctors in repairing severe bone fractures.
                          • Create prototype devices to aid in the healing of a bone fractures and test them for strength.
                          • Evaluate the strengths and weaknesses of a prototype medical device based on model testing.

                          Educational Standards

                          Each TeachEngineering lesson or activity is correlated to one or more K-12 science, technology, engineering or math (STEM) educational standards.

                          All 100,000+ K-12 STEM standards covered in TeachEngineering are collected, maintained and packaged by the Achievement Standards Network (ASN), a project of D2L (

                          In the ASN, standards are hierarchically structured: first by source e.g., by state within source by type e.g., science or mathematics within type by subtype, then by grade, etc.

                          NGSS: Next Generation Science Standards - Science

                          HS-ETS1-2. Design a solution to a complex real-world problem by breaking it down into smaller, more manageable problems that can be solved through engineering. (Grades 9 - 12)

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                          International Technology and Engineering Educators Association - Technology
                          • Medical technologies include prevention and rehabilitation, vaccines and pharmaceuticals, medical and surgical procedures, genetic engineering, and the systems within which health is protected and maintained. (Grades 9 - 12) More Details

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                          State Standards
                          Colorado - Math
                          • Quantitative reasoning is used to make sense of quantities and their relationships in problem situations. (Grades 9 - 12) More Details

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                          Colorado - Science
                          • Discuss how two or more body systems interact to promote health for the whole organism (Grades 9 - 12) More Details

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                          Materials List

                          For the teacher's introductory presentation:

                          • 1 turkey femur (drumstick)
                          • safety glasses or goggles, one per student
                          • other supplies, depending on group design (see below) , one per person

                          For the entire class to share:

                          [Note: These supplies depend on student designs, so wait to purchase them after designs are finalized.]

                          [Use any of these items that are accessible a machine shop may have some items]

                          • drill (a drill press is preferred, but a hand drill is okay)
                          • hack saw
                          • screw driver
                          • (optional) tile drill bit (makes drilling into bone easier and less likely to crack)

                          Worksheets and Attachments

                          More Curriculum Like This

                          Students learn about the role engineers and engineering play in repairing severe bone fractures. They acquire knowledge about the design and development of implant rods, pins, plates, screws and bone grafts.

                          Students learn how forces affect the human skeletal system through fractures and why certain bones are more likely to break than others depending on their design and use in the body. They learn how engineers and doctors collaborate to design effective treatments with consideration for the location, .

                          Students learn about the strength of bones and methods of helping to mend fractured bones. Working as biomedical engineers, student teams design their own splint or cast to help repair a fractured bone, learning about the strength of materials used.

                          Students are introduced to how engineering closely relates to the field of biomechanics and how the muscular system produces human movement. They learn the importance of the muscular system in our daily lives, why it is important to be able to repair muscular injuries and how engineering helps us by.

                          Pre-Req Knowledge

                          A basic understanding of bones, how they work and what they are made of. See the Our Amazing Skeleton lesson.


                          (Have ready to show to the class the attached Bone Repair Challenge PowerPoint presentation.)

                          Who has ever broken a bone? How did you repair it? When a bone breaks, it immediately begins healing itself. Usually, a doctor can assist a minor bone fracture by "immobilizing" the broken region with a cast or a sling to minimize its movement while healing. However, when severe fractures occur, sometimes more intense measures must be taken. For severe fractures, doctors must consider the risk of infection, the length of time needed to heal the break, and how to best heal the bone correctly to restore function and mobility.

                          For severe fractures, biomedical and materials engineers assist doctors by developing various devices used to help heal bones. Two categories of bone repair are internal and external fixation. Internal fixation is a temporary or permanent fixture that directly attaches to the bone under the skin for alignment and support. These include pins, rods, plates, screws, wires and bone grafting. External fixation is a temporary repair support outside of the skin that stabilizes and aligns the bone while the body heals. These devices include screws, metal braces and casts. External fixation devices can be adjusted outside of the bone. In some cases, internal fixation methods are chosen because they can provide increased patient mobility and quicker healing time.

                          Biomedical and materials engineers must consider the strength and biocompatibility of the device as well as ease of implantation and minimal invasiveness for the patient. Over the next few class periods, we will break turkey femurs and then work in groups to engineer ways to repair the bones. Let's see if you can make the bone stronger than before it was broken!

                          (Show the class the attached Bone Repair Challenge PowerPoint presentation to introduce or review the kinds of broken bones and the current medical internal fixation approaches to repair them [pins, rods, plates, screws, etc.]. The presentation includes medical illustration and x-ray examples, and concludes with the activity design challenge on one slide.)


                          • Purchase enough turkey drumsticks to equal to the number of groups plus one or two extras, the bigger the bones the better. Ask if a butcher or meat plant might donate them. Eat the turkey or remove the meat from the bones.
                          • To make the turkey femurs as clean as possible, boil them and remove any remaining meat and other tissue. If necessary, soak the bones in a solution of 90% warm water and 10% bleach or ammonia to make cleaning them easier.
                          • Let the bones dry for

                          1. Divide the class into groups of three students each.
                          2. Break the turkey femurs, keeping track of the maximum weight each bone could bear before breaking. Two suggested methods:
                          • Use a stress tester, such as an Instron universal testing machine, to break bone from the side and/or from compression. Universities often have stress testing equipment.
                          • Bridge a bone across two desk edges and hang enough weight from the center of the bone until it breaks. Expect a turkey femur to bear up to 200 lbs (91 kg), depending on its size. (This approach is described in more detail as part of the Sticks and Stones Will Break That Bone! activity.)

                          Students work together to repair a broken turkey bone.

                          1. Challenge teams to design and build repairs for their fractured bones. Encourage them to try to make the bone even stronger than before. Have students follow along with the design worksheet during this process.
                          2. Have students carefully examine the extent and nature of the bone fracture(s), and brainstorm possible ways to repair their broken turkey bones. Using an Instron universal testing machine to test the strength of a repaired turkey bone.

                          1. Have each group predict the performance of their repaired bone.
                          2. Break each reinforced bone using the same method used before.
                          3. Have students record how well their bone resisted the weight compared to its unbroken state. Record how much weight the reinforced bone withstood, and any other observations during the test. Have students complete their worksheets while others complete the stress testing.
                          4. When testing is complete, discuss and compare all results as a class.
                          5. Have students give final presentations, answering questions as described in the Assessment section.


                          biocompatibility: A characteristic of some materials that when they are inserted into the body do not produce a significant rejection or immune response.

                          bone graft: Bone taken from a patient during surgery or a bone substitute that is used to take the place of removed bone or to fill a bony defect.

                          external fixation: The process of installing temporary repair supports outside of the skin to stabilize and align bone while the body heals. Examples: screws in bone, metal braces, casts, slings.

                          fracture: An injury to a bone in which the tissue of the bone is broken.

                          internal fixation: The process of fastening together pieces of bone in a fixed position for alignment and support, using pins, rods, plates, screws, wires, grafting, and other devices, all under the skin. Can be temporary or permanent fixtures.

                          prototype: An original, full-scale, and usually working model of a new product, or new version of an existing product.


                          Brainstorming: In small groups, have students engage in open discussion. Remind them that no idea or suggestion is "silly." Respectfully listen to all ideas. Ask the students:

                          Activity Embedded Assessment

                          Design Presentations: After creating two or three design solutions for fixing the broken bone, have each group present their best design and answer the following questions:

                          1. How does the design support the weight and movement of the patient?
                          2. Is it minimally invasive (easy for a doctor to implant)? Why or why not?
                          3. Are the materials biocompatible?
                          4. Is it realistic?
                          5. What are the design strong points and weaknesses?
                          6. Which design did you choose? Why?

                          Final Presentations and Project Reflection: After testing their devices, have students consider again the questions from the activity-embedded assessment, as well as the following:

                          1. How did your repair handle the load during testing?
                          2. Where on the bone did the repair fail? Why do you think it failed there?
                          3. How could you have improved your device?

                          Safety Issues

                          • Have students wear eye protection throughout the activity as bone fragments may splinter and fly.
                          • Provide proper training and safety measures when using any power tools.

                          Troubleshooting Tips

                          Experiment with bone breaking in advance of the activity to make sure your method works well.

                          Have on hand one or two extra bones in case students have problem when fixing their original bone or with which students can practice drilling.

                          Activity Extensions

                          Have students, individually or in groups, draw final designs based on what they learned from the testing and presentations.

                          Activity Scaling

                          • For lower grades, have students work on smaller bones, such as chicken wings. Or conduct the fourth-grade Sticks and Stones Will Break That Bone! activity, which includes a class demonstration to break a chicken bone by applying a load until it fails (fractures), followed by student teams acting as biomedical engineers, designing (on paper) their own splints or casts to help mend fractured bones.
                          • For lower grades, have students repair their bones solely with external fixation, such as bracing, casts or splints.


                          Bone fracture repair-series, Procedure. Last updated September 21, 2009. MedlinePlus Medical Encyclopedia, US National Library of Medicine, National Institutes of Health. Accessed October 29, 2009.

                          Prototype. The American Heritage® Dictionary of the English Language, Fourth Edition. Houghton Mifflin Company. Accessed November 2, 2009, from website.



                          Supporting Program


                          This digital library content was developed by the Integrated Teaching and Learning Program under National Science Foundation GK-12 grant no. 0338326. However, these contents do not necessarily represent the policies of the National Science Foundation, and you should not assume endorsement by the federal government.

                          Current Methods for Skeletal Muscle Tissue Repair and Regeneration

                          Skeletal muscle has the capacity of regeneration after injury. However, for large volumes of muscle loss, this regeneration needs interventional support. Consequently, muscle injury provides an ongoing reconstructive and regenerative challenge in clinical work. To promote muscle repair and regeneration, different strategies have been developed within the last century and especially during the last few decades, including surgical techniques, physical therapy, biomaterials, and muscular tissue engineering as well as cell therapy. Still, there is a great need to develop new methods and materials, which promote skeletal muscle repair and functional regeneration. In this review, we give a comprehensive overview over the epidemiology of muscle tissue loss, highlight current strategies in clinical treatment, and discuss novel methods for muscle regeneration and challenges for their future clinical translation.

                          1. Introduction

                          Skeletal muscle is one of the most abundant tissues in the human body. It accounts for 40%–45% of the total body mass and is necessary for generating forces for movement [1]. Up to a certain threshold, skeletal muscle has the capability of regenerating lost tissue upon injury [2]. Beyond this threshold, the remaining muscle tissue is unable to fully regenerate its function. This loss of skeletal muscle with lasting functional impairment is defined as “volumetric muscle loss” (VML) [3–5]. It can substantially impact the quality of life of patients by significantly reducing the functionality of the locomotion system [4].

                          Frequent reasons for skeletal muscle injuries are high-energy traffic accidents, blast trauma, combat injuries, surgical and orthopedic situations (e.g., after compartment syndrome or tumor resection), or contusion injury during sports that lead to an acute muscle tissue loss [6, 7]. Approximately 35–55% of sport injuries involve muscle damage at the myofiber level [8]. Those injuries that involve 20% or more of muscle loss of the respective muscle mass need reconstructive surgical procedures [9]. Progressive muscle loss can result from metabolic disorders or inherited genetic diseases such as Duchenne muscular dystrophy, Amyotrophic Lateral Sclerosis, and pediatric Charcot-Marie-Tooth disease [10–13]. Muscle atrophy can also be a consequence of peripheral nerve injuries, chronic kidney disease, diabetes, and heart failure [14, 15]. Up to 20% loss of muscle mass can be compensated by the high adaptability and regenerative potential of skeletal muscle. Beyond this threshold functional impairment is inevitable and can lead to severe disability as well as cosmetic deformities, which is why therapeutic options are in urgent demand for these patients [4, 5, 16, 17].

                          Muscle regeneration relies on a heterogeneous population of satellite cells, interstitial cells, and blood vessels and is mainly controlled through ECM proteins and secreted factors [18, 19]. Normally muscle mass is maintained by a balance between protein synthesis and degradation [20]. In most cases of VML, the regeneration capability of skeletal muscles is impeded, because necessary regenerative elements, mainly satellite cells, perivascular stem cells, and the basal lamina, are physically removed [21, 22]. Through denervation, protein degradation pathways (the proteasomal and the autophagic-lysosomal pathways) are activated. Therefore protein degradation rates exceed protein synthesis, which contributes to the muscle atrophy accompanied by gradual decrease of muscle wet weight and muscle fiber diameters [23, 24].

                          Revascularization is typically impaired. The following ischemic conditions favor fibroblast proliferation, fibrosis, and fibrotic scar tissue formation, which leads to further degeneration of the muscle [25]. The ECM composition and extent in scar tissues affect many aspects of myogenesis, muscle function, and reinnervation [26]. It can severely constrain motion and thereby aggravate the consequences of muscle tissue loss. Also in chronic muscle loss like Duchenne muscular dystrophy, fibrosis is a major problem [27]. Here, the consistent breakdown of myofibers cannot be fully compensated by satellite cell proliferation. The following inflammatory processes lead to an altered production of extracellular matrix (ECM) and consequent development of fibrosis and scar tissue formation [27–29]. This scar formation can be reduced either by injection of, for example, 5-fluorouracil and bleomycin, which antagonizes fibroblast proliferation and neoangiogenesis or by laser therapy with release of contracture and functional improvements after 6–12 months’ treatment [30, 31]. Regeneration with regression of scar tissue and functional recovery can furthermore be optimized with fat grafting [32]. However, reducing scar formation is not enough for promoting muscle tissue repair and regeneration. This reinvigorates clinical and research efforts directed at replacing or regenerating larger volumes of muscle tissue.

                          2. Current Methods for Treating Muscle Tissue Loss in the Clinic

                          Current standard of care for VML is typically based on surgical intervention with autologous muscle graft and physical therapy. Further clinically used strategies include acupuncture and application of scaffolds.

                          2.1. Surgical Techniques

                          Surgical treatment for VML includes mainly scar tissue debridement and/or muscle transposition [33]. Autologous muscle transfer is commonly performed in a clinical situation, when there are large areas of muscle loss following trauma, tumor resection, or nerve injury, which impairs the irreplaceable motor function [34, 35]. The surgeons graft healthy muscle from a donor site unaffected by the injury to restore the lost or impaired function [36]. When no adjacent muscle is available because of high-level nerve injuries or severe trauma, autologous muscle transplantation together with neurorrhaphy, in the form of free functional muscle transfer, can be applied [37, 38]. The most popular autologous muscles are latissimus dorsi muscle and gracilis muscle. Latissimus dorsi muscle transfer has been shown to be safe and efficient for restoration of elbow flexion after injuries [34]. In the case of a synovial sarcoma affecting the right gluteus medius and minimus muscles, the function of the affected hip abduction could be fully reconstructed with a free neurovascular latissimus dorsi muscle transplantation [39]. Free gracilis muscle transfer is commonly utilized to restore elbow flexion after pan-brachial plexus injury [40]. It is also applied for muscle weakness after facial palsy or pelvic floor reconstruction [41, 42]. Although functional muscle flaps can lead to at least decent functional results, they cause substantial donor site morbidity and inadequate innervation [43]. Moreover, as many as 10% of these reconstructive surgeries result in complete graft failure due to complications such as infection and necrosis [44]. Sometimes, the source of autologous muscles for grafting is a problem, if the patient is severely injured.

                          2.2. Physical Therapy

                          Exercise has the ability to prevent a decrease of skeletal muscle mass [45]. Thus, in addition to surgical techniques, physical therapy is a noninvasive/minimally invasive way to promote muscle tissue repair and regeneration. It is especially used for rehabilitation after injuries and muscle tissue transfer, or to treat chronic muscle loss.

                          Physical rehabilitation aims at strengthening the remaining muscles. This has been shown to accelerate muscle healing/regeneration by modulating the immune response, release of growth factors, promoting vascularization, and reducing scar formation [46–48]. Functional performance of nonrepaired VML injured muscle could be significantly improved with physical rehabilitation in the form of voluntary wheel running [49]. Interventions to enhance angiogenesis including exercise and massage are potential strategies to accelerate new muscle formation in clinically transplanted muscle grafts or other surgical situations [50]. It has been reported that physical exercise can upregulate the IGF-1 signaling pathway and decrease myostatin in muscle tissue of animals and humans, thus preventing muscle atrophy [51–53].

                          Physical therapy can indeed improve muscle repair and recovery however, it is unable to facilitate substantial muscle regeneration within the defect areas in VML. In addition, patients with severe diseases or injuries are frequently unable to make consistent exercise, which limits physical therapy as a treatment for VML.

                          2.3. Acupuncture

                          Acupuncture is a branch of traditional Chinese medicine, which has been widely used to treat various diseases around the world [54–56]. Electrical acupuncture treatment has been shown to suppress myostatin expression, leading to satellite cell proliferation and skeletal muscle repair [57]. Acupuncture plus low-frequency electrical stimulation (Acu-LFES) could enhance muscle regeneration and prevent muscle loss by replicating the benefits of exercise through stimulation of muscle contraction [58]. It is suitable for some patients with severe diseases, which are unable to perform exercise frequently. Acu-LFES was shown to counteract diabetes-induced skeletal muscle atrophy by increasing IGF-1 and thereby stimulating muscle regeneration [58]. Application of Acu-LFES for the treatment of diabetic myopathy and muscle loss induced by chronic kidney disease showed good functional improvement of the muscle [58, 59]. The underlying mechanism includes activation of M2 microphages and reversing mRNA expression levels of the E3 ubiquitin ligase atrogin-1.

                          Similar to physical exercise, acupuncture improves muscle function restoration and stimulates muscle regeneration especially in patients with muscle atrophy after chronic diseases. However, there is limited success for the regeneration of large volume muscle defects after trauma or tumor resection. Furthermore, more work needs to be done to determine the optimal timing and intensity of Acu-LFES as a standard treatment for muscle atrophy.

                          2.4. Biological Scaffolds

                          Biological scaffolds composed of extracellular matrix (ECM) proteins are commonly used in regenerative medicine and in surgical procedures for tissue reconstruction and regeneration. The scaffolds can promote the repair of VML by providing a structural and biochemical framework [60]. For smaller amounts of muscle loss, several tissue-derived scaffolds have been tested in animal models and translated into the clinic for surgical application [6]. Xenogeneic extracellular matrix and autologous tissue have been utilized to restore functional muscle and simultaneously generate a biological niche for recovery [61]. A multilayered scaffold made of ECM derived from porcine intestinal submucosa has been applied for reconstruction of vastus medialis muscle in patients [16]. The patient showed marked gains in isokinetic performance 4 months after surgery and new muscle tissue at the implant site was demonstrated by computer tomography. Porcine small intestinal submucosa-extracellular matrix has also been utilized for the treatment of abdominal musculoskeletal wall defects, where it was sutured at the defect corners and subcuticularly closed with a vicryl-suture [61]. Also, porcine ECM from urinary bladder has been implanted in an attempt to treat VML in human beings [60]. Functional improvement with formation of muscle tissue was observed in three of the five human patients in this study.

                          However, allograft or xenogeneic scaffolds can still induce adverse immune response after decellularization and there might be potential risk of infectious disease transmission. Therefore, there is a clinical need to develop new strategies that can facilitate safe bigger muscle tissue repair and regeneration.

                          3. Developing Technologies for Muscle Tissue Engineering and Regeneration

                          To address remaining clinical problems and explore novel strategies for muscle tissue engineering and regeneration, new technologies have been investigated intensively. While tissue bioengineering approaches aim to construct complex muscle structures in vitro for subsequent implantation and replacement of the missing muscles, tissue regeneration approaches develop tissue-like scaffolds that can be implanted to enhance new muscle formation from remaining tissue in vivo [62]. Both approaches mainly rely on combinations of scaffolds, cells, and molecular signaling with differing focus.

                          3.1. Scaffold-Based Strategies

                          Biomaterials can provide chemical and physical cues to transplanted cells or host muscle cells to enhance their survival, promote their functional maturation, protect them from the foreign body responses, and recruit host cells and regenerate muscle tissues [63]. Biological scaffolds are used in a variety of clinical tissue engineering applications and have been studied in preclinical skeletal muscle VML injury models frequently over the last decade. They are mainly made of natural polymers, synthetic polymers, or ECM and attempt to create a microenvironment niche to favorably control the behavior of resident cells.

                          Natural polymers such as alginate, collagen, and fibrin have been utilized extensively in skeletal muscle engineering [64–66]. They possess intrinsic bioactive signaling cues to enhance cell behavior [67–69]. Alginate gels with a stiffness of 13–45 kPa were found to maximize myoblast proliferation and differentiation [70]. Freeze-dried collagen scaffolds facilitated the integration of aligned myotubes into a large muscle defect, which were capable of producing force upon electrical stimulation [71]. Collagen could also supply necessary growth factors to the wound site to increase muscle cell migration [72, 73]. Fibrin gels were reported to promote myoblast survival and differentiation into myofibers when integrated in tissues [74]. Fibrin scaffolds with microthread architecture were also shown to support the healing of VML in mouse models [75].

                          As the natural polymer only offers limited mechanical stiffness and can be easily degraded, a variety of synthetic materials have been used for skeletal muscle regeneration such as PGA, PLA, and PLGA [66, 76–78]. Myoblasts seeded onto electrospun meshes with aligned nanofiber orientation can fuse into highly aligned myotubes [78]. Furthermore, synthetic scaffolds can be easily engineered to facilitate the controlled release of growth factors for inducing muscle regeneration [75, 79]. The main disadvantages include typically poorer cell affinity compared to natural polymers and the risk of stimulation of a foreign body response by the polymer or its degradation products [79].

                          To improve regeneration of muscle tissues, the in vivo microenvironment of the scaffolds ideally would mimic native tissues and thereby facilitate remodeling of the neotissue [80]. An attractive approach for the repair of VML is therefore the transplantation of a myoinductive decellularized scaffold that attracts the cells required for myogenesis from the host. That is why muscle-derived ECM scaffolds are popularly investigated. These ECM scaffolds can fill the defect and restore morphology temporarily [17]. They can further be filled by bone-marrow derived mesenchymal stem cells (MSCs) after implantation. This enriched matrix gains more blood vessels and regenerates more myofibers than “conventional” extracellular matrix [17, 81]. Indeed, hydrogels derived from decellularized skeletal muscle matrix have been shown to enhance the proliferation of skeletal myoblasts when injected into an ischemic rat limb [82]. An alternative method could be to utilize minced skeletal muscle tissue that has not been decellularized, which has been reported to show better muscle regeneration than devitalized scaffolds [83]. Comparable to muscle-derived matrix, small intestinal submucosa-extracellular matrix can lead to contractile sheets of skeletal muscle with comparable contractile force [61]. For in vitro muscle tissue engineering, rat myoblasts have also been preconditioned on a porcine bladder acellular matrix in a bioreactor and then implanted in nude mice at a muscle defect to restore muscular tissue [80].

                          Another obstacle in muscle regeneration is the musculotendinous junction. This can be partly restored in absence of implanted cells by extracellular matrix-based platforms that have been shown to withstand half of the force of the contralateral site after complete resection in a mammalian model [80]. The newly formed muscle cells have shown better adherence to 3D polyurethane-based porous scaffolds with low stiffness and larger roughness values [84].

                          3.2. Cell-Based Strategies

                          Muscle fiber regeneration is performed by cells and consequently cell-based strategies for regeneration have been pursued [83, 85]. The cell types utilized for treating muscle loss mainly include myoblasts, satellite cells (SCs), mesoangioblasts, pericytes, and mesenchymal stem cells (MSCs) [86–88]. The most well characterized muscle stem cell is the satellite cell (SC). SCs are able to contribute extensively to the formation of new muscle fibers [86, 89]. SCs transplanted into dystrophin-deficient mdx mice yielded highly efficient regeneration of dystrophic muscle and improved muscle contractile function [90]. Unfortunately, in vitro expansion of SCs results in significant reduction of their ability to produce myofibers in vivo [91] and consequently, obtaining a sufficiently large number of fresh SCs for clinical application is impractical [92]. Myoblasts have been used for reconstructing muscle tissue defects with a variety of scaffolds [87, 93, 94]. They were shown to functionally integrate into the existing musculature of the host. Injection of a larger number of myoblasts into muscles showed promising results for the treatment of dystrophin-deficient models [95]. Also MSCs could be involved in myotube formation through heterotypic cell fusion after myogenic gene activation [88]. Mesoangioblasts and pericytes have been studied for treating muscular dystrophy, which resulted in increasing the force [96]. They have also been utilized in tissue engineered hydrogel carriers, with some reported success for promoting muscle regeneration [97].

                          Stem-cell-based therapies provide notable therapeutic benefits on reversing muscle atrophy and promoting muscle regeneration. Stem cell therapy (e.g., umbilical cord blood stem cell transplantation) showed positive results for treating Duchenne muscular dystrophy [98]. After application of stem cells, an increase of dystrophin positive muscular fibers was found. Biopsies of calf muscle showed growing myoblasts cells and muscular tubes and an improvement in arms and legs during physical examination was reported.

                          3.3. Molecular Signaling Based Strategies

                          Beside cues from the ECM, also a diversity of stimulatory and inhibitory growth factors such as IGF-1 and TGF-ß1 can drive endogenous skeletal muscle regeneration by activating and/or recruiting host stem cells [22]. They can be loaded on scaffolds for controlled delivery to the injured areas [72, 99]. Sustained delivery of VEGF, IGF-1, or SDF-1a was shown to enhance myogenesis and promote angiogenesis and muscle formation [73, 100–102]. Rapid release of hepatocyte growth factor (HGF) loaded on fibrin microthread scaffolds promoted remodeling of functional muscle tissue and enhanced the regeneration of skeletal muscle in mouse models [75]. Combination therapy of h-ADSCs and bFGF hydrogels resulted in functional recovery, revascularization, and reinnervation in lacerated muscles with minimal fibrosis [103]. Furthermore, PEDF peptide was reported to promote the regeneration of skeletal muscles [104].

                          Research into the pathogenesis of sarcopenia as one of the most frequent muscular diseases has elucidated different molecular pathways. The most promising targets include BMP and myostatin [105]. Indeed, medication with human recombinant BMP-2/7 and antimyostatin can help to reduce sarcopenic symptoms [106]. Cachexia is addressed with anamorelin, a ghrelin agonist, and selective androgen receptor modulator as well as anticytokines/myokines [107]. Another factor involved in muscle healing seems to be TGF-β. Increased TGF-β1 levels, which could be detected after the use of nonsteroidal anti-inflammatory drugs, helped to regenerate muscle tissue [108–110].

                          Spinal muscular atrophy arises from mutations in the survival motor neuron 1 (SMN1) gene, which often leads to the deficiency of the ubiquitous SMN protein [111]. Therefore, one of the most promising strategies is to increase the levels of full-length SMN [112]. Nusinersen is an antisense oligonucleotide drug developed for the treatment of spinal muscular atrophy (SMA), which has been approved by the US Food and Drug Administration (FDA) and European Medicines Agency (EMA) [113]. It can modulate the pre-mRNA splicing of the survival motor neuron 2 gene and showed significant improvement of muscle function after treatment. Clinical trials on infants showed significant mean improvements in developmental motor milestones including sitting, walking, and motor function [114].

                          3.4. Other Developing Techniques

                          The effect of heat stress on skeletal muscle regeneration was investigated in experimental rats [115]. Results showed that applying heat packs immediately after crush injury accelerated the degeneration process at the injured site, facilitated migration of macrophages, proliferation, and differentiation of satellite cells, and promoted muscle tissue regeneration.

                          Low-level laser therapy (LLLT) has also been evaluated as a therapeutic approach for stimulating muscle repair and recovery after endurance exercise training in rats [116]. Other results from the rat model suggest that it could also be an option to reduce fibrosis and myonecrosis triggered by bupivacaine and accelerate the muscle regeneration process [117]. As possible mechanisms, decreased inflammation and muscle creatine kinase levels are discussed. The combination of LLLT with platelet rich plasma (PRP) produced better results for promoting muscle regeneration after injuries compared to the isolated use of LLLT or PRP [118].

                          The effect of neuromuscular electrical stimulation (NMES) on skeletal muscle regeneration was assessed in healthy subjects. It increased the proliferation of myogenic precursor cells (MPCs) and their fusion with mature myofibers, which improved the regenerative capacity of skeletal muscle [119]. The effect on models with muscle injury or VML needs to be further investigated.

                          4. Challenges and Future Perspectives

                          4.1. Mechanical Properties of Biomaterials

                          Biomaterials for muscle tissue engineering and regeneration should persist long enough to support organized functional muscle regeneration and could be degraded gradually along with new tissue formation. The scaffolds created with natural polymers are usually associated with poor mechanical stiffness and rapid degradability, when not chemically crosslinked [120]. Synthetic polymers provide an artificial alternative with flexible mechanical properties [121, 122]. However, the use of synthetic scaffolds can be associated with side effects such as inhibition of cell migration and cell-to-cell communication [123].

                          A challenge for the near future will be to join the advantageous properties of natural and artificial polymers. Design of scaffolds combining favorable cell interaction with mechanical strength will facilitate implantation, give direct support to the tissue, and allow remodeling and therefore regeneration of the impaired tissue. Ideally these materials can then be used in combination with 3D-printing technology to tailor the scaffold based on the individual loss of muscle.

                          The mechanical and surface properties of the scaffold can be further engineered to affect the cell behavior in terms of adhesion, proliferation, migration, and differentiation [124]. If stem cells are seeded onto such scaffolds, they may therefore be guided to differentiate into different types of cells based on the scaffold properties [125, 126]. Moreover, degradation products from an ECM scaffold might contribute to the recruitment of host cells for tissue remodeling by chemoattraction [127]. Thus, better understanding of cell-scaffold interaction and development of a carrier scaffold that stimulates the niche environment for ongoing remodeling processes are further goals for future development in this area.

                          4.2. Vascularization in the Process of Regeneration

                          For engineering muscle constructs in vitro, one of the major limitations is the lack of vascularization [128]. It has been shown that myoblasts need to be within 150 μm of the supply route for oxygen and nutrients (typically vessels) to survive, proliferate, and differentiate [129]. This limits the size of constructs without a functional vascular network. Insufficient vascularization can lead to nutrient deficiencies and hypoxia deeper in the scaffolds, which results in nonuniform cell differentiation and integration, and thus decreases tissue functionality [130].

                          Also for in vivo muscle tissue regeneration facilitated by bioengineered muscle tissue constructs, the absence of immediate blood supply is one main reason for failure [131]. Complete revascularization of scaffolds by ingrowth of bed vessels into the graft can take up to 3 weeks, which significantly limits the capacity to obtain scar free tissue regeneration [132]. An inability of fast vascularization inevitably results in cell death and in the worst case loss of the tissue [133].

                          In order to solve this problem, different approaches for improved vascularization are conceivable: One way is administration of growth factors like bFGF, which can accelerate neoangiogenesis in the early stages of healing [134]. Another possibility is a coculture with endothelial cells [135]. In addition, integration of vascular networks into the bioengineered scaffold by microfluidic methods or bioprinting is expected to provide solutions in the near future [128, 136–138]. Maybe the combination of several approaches will eventually solve the current vascularization deficit of the designed tissues.

                          4.3. Innervation of Regenerated Muscles

                          A critical step for regenerating functional muscle tissue after VML injuries is achieving de novo innervation of regenerated myofibers (e.g., reestablishment of neuromuscular junctions, NMJs) otherwise, the regenerated muscle will become atrophic [139]. In all cases of autologous muscle transplantation, the force developed following direct or nerve stimulation is weaker than normal [140]. This is partially due to increased connective tissue and the failure of regeneration of some muscles. Another critical factor is the poor reinnervation at the sites of the original NMJs, which influences the force output [24]. It is unclear to what extent the innervation of the regenerated muscles can be restored. To rebuild the NHJs in newly regenerated muscle fibers, nerves need to be regenerated and new motor endplates have to be formed. The motor endplates not only confer functional control over the newly regenerated muscles, but also influence muscle fiber type, alignment, and size [141]. So far studies on the reinnervation of skeletal muscles have been limited to in vitro coculture of muscle cells and neurons [142, 143]. Those results showed better contractile force in nerve-muscle constructs and then in muscle-only constructs. However, full reestablishment of new nerves and motor endplates within new muscles has proven difficult, which needs to be further investigated.

                          4.4. Immune System Problems with Scaffolds and Cells

                          Matrix derived from both allografts and xenografts is often rejected because of host immune responses arising from antigens present in the donor tissue (e.g., Gal epitope, DNA, and damage associated molecular pattern molecules) [127, 144, 145]. They are typically processed by decellularization and/or chemical crosslinking to remove or cover antigenic molecules [146]. Specific decellularization techniques seem to alleviate some of these problems for ECM [147, 148]. However, remnant DNA within biological scaffolds after decellularization can still induce inflammatory reactions following implantation [149]. The host immune response to biological scaffolds differs among the sources of the raw materials from which the ECM is harvested, the processing steps, to the intended clinical application [127]. The cellular response to porcine SIS crosslinked with carbodiimide was shown to be predominated by a neutrophilic-type response, whereas foreign-body response associated with multinucleate giant cells was observed at the surgical site implanted with human dermis and porcine dermis. The host tissue response to porcine SIS showed organized connective tissue formation and muscle cells proliferation whereas the tissue response to human dermis was predominated by a persistent low-grade chronic inflammation with fibrous connective tissue formation, which might form an adverse environment for muscle tissue regeneration [150]. Therefore, the host immune reaction to biomaterials is a challenge that needs to be overcome by either designing materials that do not elicit such effects or modulating the adverse immune response.

                          Also for polymeric biomaterials, immunological compatibility remains a problem and limited biocompatibility sometimes causes local morbidity and chronic inflammation [108]. One reason could be that polymeric biomaterials attract multinucleated giant cells for disintegration [151].

                          Whether immune activation results in tissue regeneration or scarring is determined also by the availability of a stem or progenitor cell pool [152]. The cell source seems to be important with less immunogenicity in embryonic and adult stem cells [153]. Consequently, cells isolated from cord blood and autologous stem cells would be preferred for clinical application in such materials. Induced pluripotent stem cells (iPSCs) have a wide possible range of application as their production is relatively straight forward and they can differentiate in nearly every cell type. They might be able to overcome immunogenicity and ethical concerns. However, safety concerns for the use of iPSCs in patients currently result in very high regulatory barriers that will inhibit clinical translation for the foreseeable future [154]. The interactions between immune cells and resident cells are important in skeletal muscle regeneration. Macrophages, eosinophils, and regulatory T cells have been shown to activate satellite cells, which contribute to myofibers formation after injury [155–157]. In depth understanding of the immune reactions to both biological scaffolds and transplanted cells may provide clues to therapeutic avenues to promote muscle tissue regeneration. Study of the immunomodulation by scaffolds, materials, and cells in combination with subtle signaling might provide new strategies for enhancing muscle tissue regeneration through guided cell response.

                          5. Conclusion

                          Skeletal muscle injury or loss occurs in many clinical situations. Surgical techniques are highly developed and can provide good results for reconstructing muscle function, if all goes well. Surgery is always associated with considerable risks and high costs and even if successful, usually better function at one location is traded for impaired function at another location that is less important for the patient. Research into tissue engineering and regenerative cell therapy may overcome these problems. Tissue engineering solutions will have to combine biomimetic scaffolds which guide muscle tissue growth with growth factors, embedded supply routes, and relevant cells. These cells will have to directly improve local myogenic cell amount in injured or atrophic muscles, which can be expected to promote muscle regeneration. Such creative solutions will have to rely on a deep understanding of the regeneration process required for functional muscle regeneration (cell response to scaffolds, vascularization, myogenesis, and innervation), which will require further studies.

                          Conflicts of Interest

                          The authors declare that they have no conflicts of interest.


                          1. B. J. Kwee and D. J. Mooney, “Biomaterials for skeletal muscle tissue engineering,” Current Opinion in Biotechnology, vol. 47, pp. 16–22, 2017. View at: Publisher Site | Google Scholar
                          2. F. S. Tedesco, A. Dellavalle, J. Diaz-Manera, G. Messina, and G. Cossu, “Repairing skeletal muscle: regenerative potential of skeletal muscle stem cells,” The Journal of Clinical Investigation, vol. 120, no. 1, pp. 11–19, 2010. View at: Publisher Site | Google Scholar
                          3. X. Wu, B. T. Corona, X. Chen, and T. J. Walters, “A Standardized Rat Model of Volumetric Muscle Loss Injury for the Development of Tissue Engineering Therapies,” BioResearch Open Access, vol. 1, no. 6, pp. 280–290, 2012. View at: Publisher Site | Google Scholar
                          4. B. F. Grogan and J. R. Hsu, “Volumetric muscle loss,” American Academy of Orthopaedic Surgeon, vol. 19, pp. S35–S37, 2011. View at: Publisher Site | Google Scholar
                          5. B. T. Corona, J. C. Rivera, J. G. Owens, J. C. Wenke, and C. R. Rathbone, “Volumetric muscle loss leads to permanent disability following extremity trauma,” Journal of Rehabilitation Research and Development , vol. 52, no. 7, pp. 785–792, 2015. View at: Publisher Site | Google Scholar
                          6. B. E. Pollot and B. T. Corona, “Volumetric muscle loss,” Methods in Molecular Biology, vol. 1460, pp. 19–31, 2016. View at: Publisher Site | Google Scholar
                          7. A. J. Quintero, V. J. Wright, F. H. Fu, and J. Huard, “Stem cells for the treatment of skeletal muscle injury,” Clinics in Sports Medicine, vol. 28, no. 1, pp. 1–11, 2009. View at: Publisher Site | Google Scholar
                          8. P. Counsel and W. Breidahl, “Muscle injuries of the lower leg,” Seminars in Musculoskeletal Radiology, vol. 14, no. 2, pp. 162–175, 2010. View at: Publisher Site | Google Scholar
                          9. N. J. Turner and S. F. Badylak, “Regeneration of skeletal muscle,” Cell and Tissue Research, vol. 347, no. 3, pp. 759–774, 2012. View at: Publisher Site | Google Scholar
                          10. C. Saure, C. Caminiti, J. Weglinski, F. de Castro Perez, and S. Monges, “Energy expenditure, body composition, and prevalence of metabolic disorders in patients with Duchenne muscular dystrophy,” Diabetes & Metabolic Syndrome: Clinical Research & Reviews, 2017. View at: Publisher Site | Google Scholar
                          11. O. Pansarasa, D. Rossi, A. Berardinelli, and C. Cereda, “Amyotrophic lateral sclerosis and skeletal muscle: An update,” Molecular Neurobiology, vol. 49, no. 2, pp. 984–990, 2014. View at: Publisher Site | Google Scholar
                          12. A. Jani-Acsadi, S. Ounpuu, K. Pierz, and G. Acsadi, “Pediatric Charcot-Marie-Tooth Disease,” Pediatric Clinics of North America, vol. 62, no. 3, pp. 767–786, 2015. View at: Publisher Site | Google Scholar
                          13. E. M. Yiu and A. J. Kornberg, “Duchenne muscular dystrophy,” Journal of Paediatrics and Child Health, vol. 51, no. 8, pp. 759–764, 2015. View at: Publisher Site | Google Scholar
                          14. R. R. Kalyani, M. Corriere, and L. Ferrucci, “Age-related and disease-related muscle loss: the effect of diabetes, obesity, and other diseases,” The Lancet Diabetes & Endocrinology, vol. 2, no. 10, pp. 819–829, 2014. View at: Publisher Site | Google Scholar
                          15. S. H. Tuffaha et al., “Growth hormone therapy accelerates axonal regeneration, promotes motor reinnervation, and reduces muscle atrophy following peripheral nerve injury,” Plast Reconstr Surg, vol. 137, no. 6, pp. 1771–1780, 2016. View at: Google Scholar
                          16. V. J. Mase, J. R. Hsu, S. E. Wolf et al., “Clinical application of an acellular biologic scaffold for surgical repair of a large, traumatic quadriceps femoris muscle defect,” Orthopedics, vol. 33, no. 7, 2010. View at: Publisher Site | Google Scholar
                          17. E. K. Merritt, M. V. Cannon, D. W. Hammers et al., “Repair of traumatic skeletal muscle injury with bone-marrow-derived mesenchymal stem cells seeded on extracellular matrix,” Tissue Engineering Part A, vol. 16, no. 9, pp. 2871–2881, 2010. View at: Publisher Site | Google Scholar
                          18. A. Pannérec, G. Marazzi, and D. Sassoon, “Stem cells in the hood: The skeletal muscle niche,” Trends in Molecular Medicine, vol. 18, no. 10, pp. 599–606, 2012. View at: Publisher Site | Google Scholar
                          19. B. Trappmann, J. E. Gautrot, J. T. Connelly et al., “Extracellular-matrix tethering regulates stem-cell fate,” Nature Materials, vol. 11, no. 7, pp. 642–649, 2012. View at: Publisher Site | Google Scholar
                          20. B. S. Gordon, A. R. Kelleher, and S. R. Kimball, “Regulation of muscle protein synthesis and the effects of catabolic states,” The International Journal of Biochemistry & Cell Biology, vol. 45, no. 10, pp. 2147–2157, 2013. View at: Publisher Site | Google Scholar
                          21. S. B. P. Chargé and M. A. Rudnicki, “Cellular and molecular regulation of muscle regeneration,” Physiological Reviews, vol. 84, no. 1, pp. 209–238, 2004. View at: Publisher Site | Google Scholar
                          22. R. W. Ten Broek, S. Grefte, and J. W. Von Den Hoff, “Regulatory factors and cell populations involved in skeletal muscle regeneration,” Journal of Cellular Physiology, vol. 224, no. 1, pp. 7–16, 2010. View at: Publisher Site | Google Scholar
                          23. S. Schiaffino, K. A. Dyar, S. Ciciliot, B. Blaauw, and M. Sandri, “Mechanisms regulating skeletal muscle growth and atrophy,” FEBS Journal, vol. 280, no. 17, pp. 4294–4314, 2013. View at: Publisher Site | Google Scholar
                          24. P. Wu, A. Chawla, R. J. Spinner et al., “Key changes in denervated muscles and their impact on regeneration and reinnervation,” Neural Regeneration Research, vol. 9, no. 20, pp. 1796–1809, 2014. View at: Publisher Site | Google Scholar
                          25. A. L. Serrano and P. Muñoz-Cánoves, “Regulation and dysregulation of fibrosis in skeletal muscle,” Experimental Cell Research, vol. 316, no. 18, pp. 3050–3058, 2010. View at: Publisher Site | Google Scholar
                          26. K. Grzelkowska-Kowalczyk, “The importance of extracellular matrix in skeletal muscle development and function,” in Composition and Function of the Extracellular Matrix in the Human Body, F. Travascio, Ed., in Composition and Function of the Extracellular Matrix in the Human, 2016. View at: Google Scholar
                          27. Y. Kharraz, J. Guerra, P. Pessina, A. L. Serrano, and P. Muñoz-Cánoves, “Understanding the process of fibrosis in duchenne muscular dystrophy,” BioMed Research International, vol. 2014, Article ID 965631, 2014. View at: Publisher Site | Google Scholar
                          28. W. Klingler et al., “The role of fibrosis in Duchenne muscular dystrophy,” Acta Myol, vol. 31, no. 3, pp. 184–195, 2012. View at: Google Scholar
                          29. J. M. Grasman, M. J. Zayas, R. L. Page, and G. D. Pins, “Biomimetic scaffolds for regeneration of volumetric muscle loss in skeletal muscle injuries,” Acta Biomaterialia, vol. 25, pp. 2–15, 2015. View at: Publisher Site | Google Scholar
                          30. E. H. Taudorf, P. L. Danielsen, I. F. Paulsen et al., “Non-ablative fractional laser provides long-term improvement of mature burn scars - A randomized controlled trial with histological assessment,” Lasers in Surgery and Medicine, vol. 47, no. 2, pp. 141–147, 2015. View at: Publisher Site | Google Scholar
                          31. A. C. Krakowski, A. Goldenberg, L. F. Eichenfield, J.-P. Murray, and P. R. Shumaker, “Ablative fractional laser resurfacing helps treat restrictive pediatric scar contractures,” Pediatrics, vol. 134, no. 6, pp. e1700–e1705, 2014. View at: Publisher Site | Google Scholar
                          32. M. Byrne, M. O'Donnell, L. Fitzgerald, and O. P. Shelley, “Early experience with fat grafting as an adjunct for secondary burn reconstruction in the hand: Technique, hand function assessment and aesthetic outcomes,” Burns, vol. 42, no. 2, pp. 356–365, 2016. View at: Publisher Site | Google Scholar
                          33. M. Klinkenberg, S. Fischer, T. Kremer, F. Hernekamp, M. Lehnhardt, and A. Daigeler, “Comparison of anterolateral thigh, lateral arm, and parascapular free flaps with regard to donor-site morbidity and aesthetic and functional outcomes,” Plastic and Reconstructive Surgery, vol. 131, no. 2, pp. 293–302, 2013. View at: Publisher Site | Google Scholar
                          34. M. V. Stevanovic, V. G. Cuéllar, A. Ghiassi, and F. Sharpe, “Single-stage Reconstruction of Elbow Flexion Associated with Massive Soft-Tissue Defect Using the Latissimus Dorsi Muscle Bipolar Rotational Transfer,” Plastic and Reconstructive Surgery - Global Open, vol. 4, no. 9, p. e1066, 2016. View at: Publisher Site | Google Scholar
                          35. C. A. Makarewich and D. T. Hutchinson, “Tendon Transfers for Combined Peripheral Nerve Injuries,” Hand Clinics, vol. 32, no. 3, pp. 377–387, 2016. View at: Publisher Site | Google Scholar
                          36. A. Eckardt and K. Fokas, “Microsurgical reconstruction in the head and neck region: An 18-year experience with 500 consecutive cases,” Journal of Cranio-Maxillo-Facial Surgery, vol. 31, no. 4, pp. 197–201, 2003. View at: Publisher Site | Google Scholar
                          37. E. P. Estrella and T. D. Montales, “Functioning free muscle transfer for the restoration of elbow flexion in brachial plexus injury patients,” Injury, vol. 47, no. 11, pp. 2525–2533, 2016. View at: Publisher Site | Google Scholar
                          38. J. A. Bertelli and M. F. Ghizoni, “Nerve and Free Gracilis Muscle Transfers for Thumb and Finger Extension Reconstruction in Long-standing Tetraplegia,” Journal of Hand Surgery, vol. 41, no. 11, pp. e411–e416, 2016. View at: Publisher Site | Google Scholar
                          39. S. Barrera-Ochoa, J. M. Collado-Delfa, A. Sallent, A. Lluch, and R. Velez, “Free Neurovascular Latissimus Dorsi Muscle Transplantation for Reconstruction of Hip Abductors,” Plastic and Reconstructive Surgery - Global Open, vol. 5, no. 9, p. e1498, 2017. View at: Publisher Site | Google Scholar
                          40. A. A. Maldonado, M. F. Kircher, R. J. Spinner, A. T. Bishop, and A. Y. Shin, “Free Functioning Gracilis Muscle Transfer With and Without Simultaneous Intercostal Nerve Transfer to Musculocutaneous Nerve for Restoration of Elbow Flexion After Traumatic Adult Brachial Pan-Plexus Injury,” Journal of Hand Surgery, vol. 42, no. 4, pp. 293–293.e7, 2017. View at: Publisher Site | Google Scholar
                          41. S. M. Rozen, “Facial reanimation: basic surgical tools and creation of an effective toolbox for treating patients with facial paralysis. part a: functional muscle transfers in the long-term facial palsy patient,” Plastic and Reconstructive Surgery, vol. 139, no. 2, pp. 469–471, 2017. View at: Publisher Site | Google Scholar
                          42. C. S. Jones, J. Nowers, N. J. Smart, J. Coelho, A. Watts, and I. R. Daniels, “Pelvic floor reconstruction with bilateral gracilis flaps following extralevator abdominoperineal excision - a video vignette,” Colorectal Disease, vol. 19, no. 12, pp. 1120-1121, 2017. View at: Publisher Site | Google Scholar
                          43. C. Lin, Y. Lin, C. Chen, and F. Wei, “Free Functioning Muscle Transfer for Lower Extremity Post-Traumatic Composite Structure and Functional Defect,” Journal of Reconstructive Microsurgery, vol. 119, no. 7, pp. 2118–2126, 2007. View at: Publisher Site | Google Scholar
                          44. B. Bianchi, C. Copelli, S. Ferrari, A. Ferri, and E. Sesenna, “Free flaps: Outcomes and complications in head and neck reconstructions,” Journal of Cranio-Maxillo-Facial Surgery, vol. 37, no. 8, pp. 438–442, 2009. View at: Publisher Site | Google Scholar
                          45. X. H. Wang, J. Du, J. D. Klein, J. L. Bailey, and W. E. Mitch, “Exercise ameliorates chronic kidney disease-induced defects in muscle protein metabolism and progenitor cell function,” Kidney International, vol. 76, no. 7, pp. 751–759, 2009. View at: Publisher Site | Google Scholar
                          46. T. M. Gregory, R. A. Heckmann, and R. S. Francis, “The effect of exercise on the presence of leukocytes, erythrocytes and collagen fibers in skeletal muscle after contusion,” J Manipulative Physiol Ther, vol. 18, no. 2, pp. 72–80, 1995. View at: Google Scholar
                          47. T. D. Brutsaert et al., “Regional differences in expression of VEGF mRNA in rat gastrocnemius following 1 hr exercise or electrical stimulation,” BMC Physiol, vol. 2, p. 8, 2002. View at: Publisher Site | Google Scholar
                          48. F. E. Faria et al., “The onset and duration of mobilization affect the regeneration in the rat muscle,” Histol Histopathol, vol. 23, no. 5, pp. 565–571, 2008. View at: Google Scholar
                          49. A. Aurora, K. Garg, B. T. Corona, and T. J. Walters, “Physical rehabilitation improves muscle function following volumetric muscle loss injury,” BMC Sports Science, Medicine and Rehabilitation, vol. 6, no. 1, article no. 41, 2014. View at: Publisher Site | Google Scholar
                          50. W. Andrzejewski, K. Kassolik, C. Kobierzycki et al., “Increased skeletal muscle expression of VEGF induced by massage and exercise,” Folia Histochemica et Cytobiologica, vol. 53, no. 2, pp. 145–151, 2015. View at: Publisher Site | Google Scholar
                          51. Y. Chen, S. Sood, J. Biada, R. Roth, and R. Rabkin, “Increased workload fully activates the blunted IRS-1/PI3-kinase/Akt signaling pathway in atrophied uremic muscle,” Kidney International, vol. 73, no. 7, pp. 848–855, 2008. View at: Publisher Site | Google Scholar
                          52. K. M. Majchrzak, L. B. Pupim, P. J. Flakoll, and T. A. Ikizler, “Resistance exercise augments the acute anabolic effects of intradialytic oral nutritional supplementation,” Nephrology Dialysis Transplantation , vol. 23, no. 4, pp. 1362–1369, 2008. View at: Publisher Site | Google Scholar
                          53. J. D. Kopple, H. Wang, R. Casaburi et al., “Exercise in maintenance hemodialysis patients induces transcriptional changes in genes favoring anabolic muscle,” Journal of the American Society of Nephrology, vol. 18, no. 11, pp. 2975–2986, 2007. View at: Publisher Site | Google Scholar
                          54. K. Zhao, “Acupuncture for the treatment of insomnia,” International Review of Neurobiology, vol. 111, pp. 217–234, 2013. View at: Publisher Site | Google Scholar
                          55. N. E. Haddad and O. Palesh, “Acupuncture in the treatment of cancer-related psychological symptoms,” Integrative Cancer Therapies, vol. 13, no. 5, pp. 371–385, 2014. View at: Publisher Site | Google Scholar
                          56. M. A. Urruela and M. E. Suarez-Almazor, “Acupuncture in the treatment of rheumatic diseases,” Current Rheumatology Reports, vol. 14, no. 6, pp. 589–597, 2012. View at: Publisher Site | Google Scholar
                          57. Y. Takaoka, M. Ohta, A. Ito et al., “Electroacupuncture suppresses myostatin gene expression: Cell proliferative reaction in mouse skeletal muscle,” Physiological Genomics, vol. 30, no. 2, pp. 102–110, 2007. View at: Publisher Site | Google Scholar
                          58. Z. Su, A. Robinson, L. Hu et al., “Acupuncture plus low-frequency electrical stimulation (Acu-LFES) attenuates diabetic myopathy by enhancing muscle regeneration,” PLoS ONE, vol. 10, no. 7, Article ID e0134511, 2015. View at: Publisher Site | Google Scholar
                          59. L. Hu, J. D. Klein, F. Hassounah et al., “Low-frequency electrical stimulation attenuates muscle atrophy in CKD-a potential treatment strategy,” Journal of the American Society of Nephrology, vol. 26, no. 3, pp. 626–635, 2015. View at: Publisher Site | Google Scholar
                          60. B. M. Sicari, J. P. Rubin, C. L. Dearth et al., “An acellular biologic scaffold promotes skeletal muscle formation in mice and humans with volumetric muscle loss,” Science Translational Medicine, vol. 6, no. 234, Article ID 234ra58, 2014. View at: Publisher Site | Google Scholar
                          61. J. E. Valentin, N. J. Turner, T. W. Gilbert, and S. F. Badylak, “Functional skeletal muscle formation with a biologic scaffold,” Biomaterials, vol. 31, no. 29, pp. 7475–7484, 2010. View at: Publisher Site | Google Scholar
                          62. L. Wang, L. Cao, J. Shansky, Z. Wang, D. Mooney, and H. Vandenburgh, “Minimally invasive approach to the repair of injured skeletal muscle with a shape-memory scaffold,” Molecular Therapy, vol. 22, no. 8, pp. 1441–1449, 2014. View at: Publisher Site | Google Scholar
                          63. C. A. Cezar and D. J. Mooney, “Biomaterial-based delivery for skeletal muscle repair,” Advanced Drug Delivery Reviews, vol. 84, pp. 188–197, 2015. View at: Publisher Site | Google Scholar
                          64. E. Hill, T. Boontheekul, and D. J. Mooney, “Regulating activation of transplanted cells controls tissue regeneration,” Proceedings of the National Acadamy of Sciences of the United States of America, vol. 103, no. 8, pp. 2494–2499, 2006. View at: Publisher Site | Google Scholar
                          65. D. G. Moon, G. Christ, J. D. Stitzel, A. Atala, and J. J. Yoo, “Cyclic mechanical preconditioning improves engineered muscle contraction,” Tissue Engineering - Part A., vol. 14, no. 4, pp. 473–482, 2008. View at: Publisher Site | Google Scholar
                          66. A. Lesman, J. Koffler, R. Atlas, Y. J. Blinder, Z. Kam, and S. Levenberg, “Engineering vessel-like networks within multicellular fibrin-based constructs,” Biomaterials, vol. 32, no. 31, pp. 7856–7869, 2011. View at: Publisher Site | Google Scholar
                          67. S. J. Bidarra, C. C. Barrias, and P. L. Granja, “Injectable alginate hydrogels for cell delivery in tissue engineering,” Acta Biomaterialia, vol. 10, no. 4, pp. 1646–1662, 2014. View at: Publisher Site | Google Scholar
                          68. B. D. Walters and J. P. Stegemann, “Strategies for directing the structure and function of three-dimensional collagen biomaterials across length scales,” Acta Biomaterialia, vol. 10, no. 4, pp. 1488–1501, 2014. View at: Publisher Site | Google Scholar
                          69. A. C. Brown and T. H. Barker, “Fibrin-based biomaterials: modulation of macroscopic properties through rational design at the molecular level,” Acta Biomaterialia, vol. 10, no. 4, pp. 1502–1514, 2014. View at: Publisher Site | Google Scholar
                          70. T. Boontheekul, E. E. Hill, H.-J. Kong, and D. J. Mooney, “Regulating myoblast phenotype through controlled gel stiffness and degradation,” Tissue Engineering Part A, vol. 13, no. 7, pp. 1431–1442, 2007. View at: Publisher Site | Google Scholar
                          71. V. Kroehne, I. Heschel, F. Schügner, D. Lasrich, J. W. Bartsch, and H. Jockusch, “Use of a novel collagen matrix with oriented pore structure for muscle cell differentiation in cell culture and in grafts,” Journal of Cellular and Molecular Medicine, vol. 12, no. 5A, pp. 1640–1648, 2008. View at: Publisher Site | Google Scholar
                          72. S. P. Frey, H. Jansen, M. J. Raschke, R. H. Meffert, and S. Ochman, “VEGF improves skeletal muscle regeneration after acute trauma and reconstruction of the limb in a rabbit model,” Clinical Orthopaedics and Related Research, vol. 470, no. 12, pp. 3607–3614, 2012. View at: Publisher Site | Google Scholar
                          73. Y. M. Ju, A. Atala, J. J. Yoo, and S. J. Lee, “In situ regeneration of skeletal muscle tissue through host cell recruitment,” Acta Biomaterialia, vol. 10, no. 10, pp. 4332–4339, 2014. View at: Publisher Site | Google Scholar
                          74. J. P. Beier, U. Kneser, J. Stern-Sträter, G. B. Stark, and A. D. Bach, “Y Chromosome Detection of Three-Dimensional Tissue-Engineered Skeletal Muscle Constructs in a Syngeneic Rat Animal Model,” Cell Transplantation, vol. 13, no. 1, pp. 45–53, 2004. View at: Publisher Site | Google Scholar
                          75. J. M. Grasman, D. M. Do, R. L. Page, and G. D. Pins, “Rapid release of growth factors regenerates force output in volumetric muscle loss injuries,” Biomaterials, vol. 72, pp. 49–60, 2015. View at: Publisher Site | Google Scholar
                          76. E. M. Cronin, F. A. Thurmond, R. Bassel-Duby et al., “Protein-coated poly(L-lactic acid) fibers provide a substrate for differentiation of human skeletal muscle cells,” Journal of Biomedical Materials Research Part B: Applied Biomaterials, vol. 69A, no. 3, pp. 373–381, 2004. View at: Publisher Site | Google Scholar
                          77. M. E. Hoque, W. Y. San, F. Wei et al., “Processing of polycaprolactone and polycaprolactone-based copolymers into 3D scaffolds, and their cellular responses,” Tissue Engineering Part: A, vol. 15, no. 10, pp. 3013–3024, 2009. View at: Publisher Site | Google Scholar
                          78. A. G. Guex, D. L. Birrer, G. Fortunato, H. T. Tevaearai, and M.-N. Giraud, “Anisotropically oriented electrospun matrices with an imprinted periodic micropattern: A new scaffold for engineered muscle constructs,” Biomedical Materials, vol. 8, no. 2, Article ID 021001, 2013. View at: Publisher Site | Google Scholar
                          79. J. Yang, B. Jao, A. K. Mcnally, and J. M. Anderson, “In vivo quantitative and qualitative assessment of foreign body giant cell formation on biomaterials in mice deficient in natural killer lymphocyte subsets, mast cells, or the interleukin-4 receptorα and in severe combined immunodeficient mice,” Journal of Biomedical Materials Research Part A, vol. 102, no. 6, pp. 2017–2023, 2014. View at: Publisher Site | Google Scholar
                          80. N. J. Turner, A. J. Yates, D. J. Weber et al., “Xenogeneic extracellular matrix as an inductive scaffold for regeneration of a functioning musculotendinous junction,” Tissue Engineering Part A, vol. 16, no. 11, pp. 3309–3317, 2010. View at: Publisher Site | Google Scholar
                          81. E. K. Merritt, D. W. Hammers, M. Tierney, L. J. Suggs, T. J. Walters, and R. P. Farrar, “Functional assessment of skeletal muscle regeneration utilizing homologous extracellular matrix as scaffolding,” Tissue Engineering Part A, vol. 16, no. 4, pp. 1395–1405, 2010. View at: Publisher Site | Google Scholar
                          82. J. A. DeQuach, J. E. Lin, C. Cam et al., “Injectable skeletal muscle matrix hydrogel promotes neovascularization and muscle cell infiltration in a hindlimb ischemia model,” European Cells and Materials, vol. 23, pp. 400–412, 2012. View at: Publisher Site | Google Scholar
                          83. K. Garg, C. L. Ward, C. R. Rathbone, and B. T. Corona, “Transplantation of devitalized muscle scaffolds is insufficient for appreciable de novo muscle fiber regeneration after volumetric muscle loss injury,” Cell and Tissue Research, vol. 358, no. 3, pp. 857–873, 2014. View at: Publisher Site | Google Scholar
                          84. L. Vannozzi, L. Ricotti, T. Santaniello et al., “3D porous polyurethanes featured by different mechanical properties: Characterization and interaction with skeletal muscle cells,” Journal of the Mechanical Behavior of Biomedical Materials, vol. 75, pp. 147–159, 2017. View at: Publisher Site | Google Scholar
                          85. B. N. Brown, J. E. Valentin, A. M. Stewart-Akers, G. P. McCabe, and S. F. Badylak, “Macrophage phenotype and remodeling outcomes in response to biologic scaffolds with and without a cellular component,” Biomaterials, vol. 30, no. 8, pp. 1482–1491, 2009. View at: Publisher Site | Google Scholar
                          86. C. A. Collins, I. Olsen, P. S. Zammit et al., “Stem cell function, self-renewal, and behavioral heterogeneity of cells from the adult muscle satellite cell niche,” Cell, vol. 122, no. 2, pp. 289–301, 2005. View at: Publisher Site | Google Scholar
                          87. M. A. MacHingal, B. T. Corona, T. J. Walters et al., “A tissue-engineered muscle repair construct for functional restoration of an irrecoverable muscle injury in a murine model,” Tissue Engineering Part A, vol. 17, no. 17-18, pp. 2291–2303, 2011. View at: Publisher Site | Google Scholar
                          88. J.-H. Lee, P. A. Kosinski, and D. M. Kemp, “Contribution of human bone marrow stem cells to individual skeletal myotubes followed by myogenic gene activation,” Experimental Cell Research, vol. 307, no. 1, pp. 174–182, 2005. View at: Publisher Site | Google Scholar
                          89. A. Sacco, F. Mourkioti, R. Tran et al., “Short telomeres and stem cell exhaustion model duchenne muscular dystrophy in mdx/mTR mice,” Cell, vol. 143, no. 7, pp. 1059–1071, 2010. View at: Publisher Site | Google Scholar
                          90. M. Cerletti, S. Jurga, C. A. Witczak et al., “Highly efficient, functional engraftment of skeletal muscle stem cells in dystrophic muscles,” Cell, vol. 134, no. 1, pp. 37–47, 2008. View at: Publisher Site | Google Scholar
                          91. D. Montarras, J. Morgan, C. Colins et al., “Developmental biology: direct isolation of satellite cells for skeletal muscle regeneration,” Science, vol. 309, no. 5743, pp. 2064–2067, 2005. View at: Publisher Site | Google Scholar
                          92. J. Meng, F. Muntoni, and J. E. Morgan, “Stem cells to treat muscular dystrophies - Where are we?” Neuromuscular Disorders, vol. 21, no. 1, pp. 4–12, 2011. View at: Publisher Site | Google Scholar
                          93. C. Borselli, C. A. Cezar, D. Shvartsman, H. H. Vandenburgh, and D. J. Mooney, “The role of multifunctional delivery scaffold in the ability of cultured myoblasts to promote muscle regeneration,” Biomaterials, vol. 32, no. 34, pp. 8905–8914, 2011. View at: Publisher Site | Google Scholar
                          94. M. T. Wolf, K. A. Daly, J. E. Reing, and S. F. Badylak, “Biologic scaffold composed of skeletal muscle extracellular matrix,” Biomaterials, vol. 33, no. 10, pp. 2916–2925, 2012. View at: Publisher Site | Google Scholar
                          95. R. Miller, K. Sharma, G. Pavlath et al., “Myoblast implantation in Duchenne muscular dystrophy: The San Francisco study,” Muscle & Nerve, vol. 20, no. 4, pp. 469–478. View at: Publisher Site | Google Scholar
                          96. M. Sampaolesi, S. Blot, G. D'Antona et al., “Mesoangioblast stem cells ameliorate muscle function in dystrophic dogs,” Nature, vol. 444, no. 7119, pp. 574–579, 2006. View at: Publisher Site | Google Scholar
                          97. C. Fuoco, M. Salvatori, A. Biondo et al., “Injectable polyethylene glycol-fibrinogen hydrogel adjuvant improves survival and differentiation of transplanted mesoangioblasts in acute and chronic skeletal-muscle degeneration,” Skeletal Muscle, vol. 2, no. 1, p. 24, 2012. View at: Publisher Site | Google Scholar
                          98. C. Zhang et al., “Therapy of Duchenne muscular dystrophy with umbilical cord blood stem cell transplantation,” Zhonghua Yi Xue Yi Chuan Xue Za Zhi, vol. 22, no. 4, pp. 399–405, 2005. View at: Google Scholar
                          99. D. W. Hammers, A. Sarathy, C. B. Pham, C. T. Drinnan, R. P. Farrar, and L. J. Suggs, “Controlled release of IGF-I from a biodegradable matrix improves functional recovery of skeletal muscle from ischemia/reperfusion,” Biotechnology and Bioengineering, vol. 109, no. 4, pp. 1051–1059, 2012. View at: Publisher Site | Google Scholar
                          100. C. Borselli, H. Storrie, F. Benesch-Lee et al., “Functional muscle regeneration with combined delivery of angiogenesis and myogenesis factors,” Proceedings of the National Acadamy of Sciences of the United States of America, vol. 107, no. 8, pp. 3287–3292, 2010. View at: Publisher Site | Google Scholar
                          101. D. Shvartsman, H. Storrie-White, K. Lee et al., “Sustained delivery of VEGF maintains innervation and promotes reperfusion in ischemic skeletal muscles via NGF/GDNF signaling,” Molecular Therapy, vol. 22, no. 7, pp. 1243–1253, 2014. View at: Publisher Site | Google Scholar
                          102. V. Y. Rybalko, C. B. Pham, P.-L. Hsieh et al., “Controlled delivery of SDF-1α and IGF-1: CXCR4+ cell recruitment and functional skeletal muscle recovery,” Biomaterials Science, vol. 3, no. 11, pp. 1475–1486, 2015. View at: Publisher Site | Google Scholar
                          103. J. H. Hwang, I. G. Kim, S. Piao et al., “Combination therapy of human adipose-derived stem cells and basic fibroblast growth factor hydrogel in muscle regeneration,” Biomaterials, vol. 34, no. 25, pp. 6037–6045, 2013. View at: Publisher Site | Google Scholar
                          104. T.-C. Ho, Y.-P. Chiang, C.-K. Chuang et al., “PEDF-derived peptide promotes skeletal muscle regeneration through its mitogenic effect on muscle progenitor cells,” American Journal of Physiology-Cell Physiology, vol. 309, no. 3, pp. C159–C168, 2015. View at: Publisher Site | Google Scholar
                          105. S. A. Saul D and R. L. Kosinsky, “Why age matters: inflammation, cancer and hormones in the development of sarcopenia,” Journal of Osteoporosis and Physical Activity, vol. 5, no. 1, 2017. View at: Publisher Site | Google Scholar
                          106. M. Scimeca, E. Piccirilli, F. Mastrangeli et al., “Bone Morphogenetic Proteins and myostatin pathways: Key mediator of human sarcopenia,” Journal of Translational Medicine, vol. 15, no. 1, article no. 34, 2017. View at: Publisher Site | Google Scholar
                          107. A. Molfino, M. I. Amabile, F. Rossi Fanelli, and M. Muscaritoli, “Novel therapeutic options for cachexia and sarcopenia,” Expert Opinion on Biological Therapy, vol. 16, no. 10, pp. 1239–1244, 2016. View at: Publisher Site | Google Scholar
                          108. R. Berebichez-Fridman, R. Gómez-García, J. Granados-Montiel et al., “The Holy Grail of Orthopedic Surgery: Mesenchymal Stem Cells - Their Current Uses and Potential Applications,” Stem Cells International, vol. 2017, Article ID 2638305, 2017. View at: Publisher Site | Google Scholar
                          109. S. S. Tseng, M. A. Lee, and A. H. Reddi, “Nonunions and the potential of stem cells in fracture-healing,” The Journal of Bone & Joint Surgery—American Volume, vol. 90, supplement 1, pp. 92–98, 2008. View at: Publisher Site | Google Scholar
                          110. Z. Qu-Petersen, B. Deasy, R. Jankowski et al., “Identification of a novel population of muscle stem cells in mice: potential for muscle regeneration,” The Journal of Cell Biology, vol. 157, no. 5, pp. 851–864, 2002. View at: Publisher Site | Google Scholar
                          111. U. R. Monani, “Spinal muscular atrophy: A deficiency in a ubiquitous protein a motor neuron-specific disease,” Neuron, vol. 48, no. 6, pp. 885–896, 2005. View at: Publisher Site | Google Scholar
                          112. V. Parente and S. Corti, “Advances in spinal muscular atrophy therapeutics,” Therapeutic Advances in Neurological Disorders, vol. 11, p. 175628561875450, 2018. View at: Publisher Site | Google Scholar
                          113. E. Mercuri, B. T. Darras, C. A. Chiriboga et al., “Nusinersen versus Sham Control in Later-Onset Spinal Muscular Atrophy,” The New England Journal of Medicine, vol. 378, no. 7, pp. 625–635, 2018. View at: Publisher Site | Google Scholar
                          114. R. S. Finkel, C. A. Chiriboga, J. Vajsar et al., “Treatment of infantile-onset spinal muscular atrophy with nusinersen: a phase 2, open-label, dose-escalation study,” The Lancet, vol. 388, no. 10063, pp. 3017–3026, 2016. View at: Publisher Site | Google Scholar
                          115. K. Takeuchi, T. Hatade, S. Wakamiya, N. Fujita, T. Arakawa, and A. Miki, “Heat stress promotes skeletal muscle regeneration after crush injury in rats,” Acta Histochemica, vol. 116, no. 2, pp. 327–334, 2014. View at: Publisher Site | Google Scholar
                          116. L. Assis, F. Yamashita, A. M. P. Magri, K. R. Fernandes, L. Yamauchi, and A. C. M. Renno, “Effect of low-level laser therapy (808 nm) on skeletal muscle after endurance exercise training in rats,” Brazilian Journal of Physical Therapy, vol. 19, no. 6, pp. 457–465, 2015. View at: Publisher Site | Google Scholar
                          117. C. N. Alessi Pissulin, A. A. Henrique Fernandes, A. M. Sanchez Orellana, R. C. Rossi e Silva, and S. M. Michelin Matheus, “Low-level laser therapy (LLLT) accelerates the sternomastoid muscle regeneration process after myonecrosis due to bupivacaine,” Journal of Photochemistry and Photobiology B: Biology, vol. 168, pp. 30–39, 2017. View at: Publisher Site | Google Scholar
                          118. T. A. Garcia, R. C. Camargo, T. E. Koike, G. A. Ozaki, R. C. Castoldi, and J. C. Camargo Filho, “Histological analysis of the association of low level laser therapy and platelet-rich plasma in regeneration of muscle injury in rats,” Brazilian Journal of Physical Therapy, vol. 21, no. 6, pp. 425–433, 2017. View at: Publisher Site | Google Scholar
                          119. E. S. D. Filippo, R. Mancinelli, M. Marrone et al., “Neuromuscular electrical stimulation improves skeletal muscle regeneration through satellite cell fusion with myofibers in healthy elderly subjects,” Journal of Applied Physiology, vol. 123, no. 3, pp. 501–512, 2017. View at: Publisher Site | Google Scholar
                          120. A. A. Chaudhari, K. Vig, D. R. Baganizi et al., “Future prospects for scaffolding methods and biomaterials in skin tissue engineering: a review,” International Journal of Molecular Sciences, vol. 17, no. 12, article no. 1974, 2016. View at: Publisher Site | Google Scholar
                          121. J. Liu, H. Zheng, P. Poh, H. Machens, and A. Schilling, “Hydrogels for Engineering of Perfusable Vascular Networks,” International Journal of Molecular Sciences, vol. 16, no. 7, pp. 15997–16016, 2015. View at: Publisher Site | Google Scholar
                          122. B. Guo and P. X. Ma, “Synthetic biodegradable functional polymers for tissue engineering: a brief review,” SCIENCE CHINA Chemistry, vol. 57, no. 4, pp. 490–500, 2014. View at: Publisher Site | Google Scholar
                          123. J. E. Reing, L. Zhang, J. Myers-Irvin et al., “Degradation products of extracellular matrix affect cell migration and proliferation,” Tissue Engineering Part A, vol. 15, no. 3, pp. 605–614, 2009. View at: Publisher Site | Google Scholar
                          124. M. Griffin, L. Nayyer, P. E. Butler, R. G. Palgrave, A. M. Seifalian, and D. M. Kalaskar, “Development of mechano-responsive polymeric scaffolds using functionalized silica nano-fillers for the control of cellular functions,” Nanomedicine: Nanotechnology, Biology and Medicine, vol. 12, no. 6, pp. 1725–1733, 2016. View at: Publisher Site | Google Scholar
                          125. E. K. F. Yim and M. P. Sheetz, “Force-dependent cell signaling in stem cell differentiation,” Stem Cell Research & Therapy, vol. 3, no. 5, article no. 41, 2012. View at: Publisher Site | Google Scholar
                          126. S. Ghassemi, G. Meacci, S. Liu et al., “Cells test substrate rigidity by local contractions on submicrometer pillars,” Proceedings of the National Acadamy of Sciences of the United States of America, vol. 109, no. 14, pp. 5328–5333, 2012. View at: Publisher Site | Google Scholar
                          127. S. F. Badylak and T. W. Gilbert, “Immune response to biologic scaffold materials,” Seminars in Immunology, vol. 20, no. 2, pp. 109–116, 2008. View at: Publisher Site | Google Scholar
                          128. J. Liu, H. Zheng, F. Krempl, L. Su, H.-G. Machens, and A. F. Schilling, “Open source 3D-printing approach for economic and fast engineering of perfusable vessel-like channels within cell-laden hydrogels,” 3D Printing and Additive Manufacturing, vol. 3, no. 1, pp. 23–31, 2016. View at: Publisher Site | Google Scholar
                          129. R. G. Dennis and P. E. Kosnik II, “Excitability and isometric contractile properties of mammalian skeletal muscle constructs engineered in vitro,” In Vitro Cellular & Developmental Biology - Animal, vol. 36, no. 5, pp. 327–335, 2000. View at: Publisher Site | Google Scholar
                          130. C.-H. Lee, S.-H. Chang, W.-J. Chen et al., “Augmentation of diabetic wound healing and enhancement of collagen content using nanofibrous glucophage-loaded collagen/PLGA scaffold membranes,” Journal of Colloid and Interface Science, vol. 439, pp. 88–97, 2015. View at: Publisher Site | Google Scholar
                          131. H. Bi and Y. Jin, “Current progress of skin tissue engineering: Seed cells, bioscaffolds, and construction strategies,” Burns & Trauma, vol. 1, no. 2, pp. 63–72, 2013. View at: Publisher Site | Google Scholar
                          132. T. T. Nyame, H. A. Chiang, T. Leavitt, M. Ozambela, and D. P. Orgill, “Tissue-Engineered Skin Substitutes,” Plastic and Reconstructive Surgery, vol. 136, no. 6, pp. 1379–1388, 2015. View at: Publisher Site | Google Scholar
                          133. M. Varkey, J. Ding, and E. Tredget, “Advances in skin substitutes—potential of tissue engineered skin for facilitating anti-fibrotic healing,” Journal of Functional Biomaterials, vol. 6, no. 3, pp. 547–563, 2015. View at: Publisher Site | Google Scholar
                          134. S. Thomopoulos et al., “The effects of exogenous basic fibroblast growth factor on intrasynovial flexor tendon healing in a canine model,” J Bone Joint Surg Am, vol. 92, no. 13, pp. 2285–2293, 2010. View at: Google Scholar
                          135. H. Haro, T. Kato, H. Komori, M. Osada, and K. Shinomiya, “Vascular endothelial growth factor (VEGF)-induced angiogenesis in herniated disc resorption,” Journal of Orthopaedic Research, vol. 20, no. 3, pp. 409–415, 2002. View at: Publisher Site | Google Scholar
                          136. Y. Nashimoto, T. Hayashi, I. Kunita et al., “Integrating perfusable vascular networks with a three-dimensional tissue in a microfluidic device,” Integrative Biology, vol. 9, no. 6, pp. 506–518, 2017. View at: Publisher Site | Google Scholar
                          137. H. Gudapati, M. Dey, and I. Ozbolat, “A comprehensive review on droplet-based bioprinting: Past, present and future,” Biomaterials, vol. 102, pp. 20–42, 2016. View at: Publisher Site | Google Scholar
                          138. K. A. DiVito, M. A. Daniele, S. A. Roberts, F. S. Ligler, and A. A. Adams, “Microfabricated blood vessels undergo neoangiogenesis,” Biomaterials, vol. 138, pp. 142–152, 2017. View at: Publisher Site | Google Scholar
                          139. T. A. Järvinen, T. L. Järvinen, M. Kääriäinen et al., “Muscle injuries: optimising recovery,” Best Practice & Research Clinical Rheumatology, vol. 21, no. 2, pp. 317–331, 2007. View at: Publisher Site | Google Scholar
                          140. I. Degreef, P. Debeer, B. Van Herck, E. Van Den Eeden, K. Peers, and L. De Smet, “Treatment of irreparable rotator cuff tears by latissimus dorsi muscle transfer,” Acta Orthopædica Belgica, vol. 71, no. 6, pp. 667–671, 2005. View at: Google Scholar
                          141. D. Chen, S. Chen, W. Wang et al., “Functional modulation of satellite cells in long-term denervated human laryngeal muscle,” The Laryngoscope, vol. 120, no. 2, pp. 353–358, 2010. View at: Publisher Site | Google Scholar
                          142. L. M. Larkin, J. H. Van Der Meulen, R. G. Dennis, and J. B. Kennedy, “Functional evaluation of nerve-skeletal muscle constructs engineered in vitro,” In Vitro Cellular & Developmental Biology - Animal, vol. 42, no. 3-4, pp. 75–82, 2006. View at: Publisher Site | Google Scholar
                          143. M. Das, J. W. Rumsey, C. A. Gregory et al., “Embryonic motoneuron-skeletal muscle co-culture in a defined system,” Neuroscience, vol. 146, no. 2, pp. 481–488, 2007. View at: Publisher Site | Google Scholar
                          144. A. Burd and T. Chiu, “Allogenic skin in the treatment of burns,” Clinics in Dermatology, vol. 23, no. 4, pp. 376–387, 2005. View at: Publisher Site | Google Scholar
                          145. M. T. Lotze, A. Deisseroth, and A. Rubartelli, “Damage associated molecular pattern molecules,” Clinical Immunology, vol. 124, no. 1, pp. 1–4, 2007. View at: Publisher Site | Google Scholar
                          146. T. W. Gilbert, T. L. Sellaro, and S. F. Badylak, “Decellularization of tissues and organs,” Biomaterials, vol. 27, no. 19, pp. 3675–3683, 2006. View at: Publisher Site | Google Scholar
                          147. M. Bottagisio, A. F. Pellegata, F. Boschetti, M. Ferroni, M. Moretti, and A. B. Lovati, “A new strategy for the decellularisation of large equine tendons as biocompatible tendon substitutes,” European Cells and Materials, vol. 32, pp. 58–73, 2016. View at: Publisher Site | Google Scholar
                          148. T. Kaully, K. Kaufman-Francis, A. Lesman, and S. Levenberg, “Vascularization—the conduit to viable engineered tissues,” Tissue Engineering Part B: Reviews, vol. 15, no. 2, pp. 159–169, 2009. View at: Publisher Site | Google Scholar
                          149. M.-H. Zheng, J. Chen, Y. Kirilak, C. Willers, J. Xu, and D. Wood, “Porcine small intestine submucosa (SIS) is not an acellular collagenous matrix and contains porcine DNA: possible implications in human implantation,” Journal of Biomedical Materials Research Part B: Applied Biomaterials, vol. 73, no. 1, pp. 61–67, 2005. View at: Publisher Site | Google Scholar
                          150. S. Badylak, K. Kokini, B. Tullius, A. Simmons-Byrd, and R. Morff, “Morphologic study of small intestinal submucosa as a body wall repair device,” Journal of Surgical Research, vol. 103, no. 2, pp. 190–202, 2002. View at: Publisher Site | Google Scholar
                          151. S. Al-Maawi, A. Orlowska, R. Sader, C. James Kirkpatrick, and S. Ghanaati, “In vivo cellular reactions to different biomaterials—Physiological and pathological aspects and their consequences,” Seminars in Immunology, vol. 29, pp. 49–61, 2017. View at: Publisher Site | Google Scholar
                          152. A. B. Aurora and E. N. Olson, “Immune modulation of stem cells and regeneration,” Cell Stem Cell, vol. 15, no. 1, pp. 14–25, 2014. View at: Publisher Site | Google Scholar
                          153. F. Bifari, “Immunological properties of embryonic and adult stem cells,” World Journal of Stem Cells, vol. 2, no. 3, pp. 50–60, 2010. View at: Publisher Site | Google Scholar
                          154. R. Yang et al., “Generation of folliculogenic human epithelial stem cells from induced pluripotent stem cells,” Nat Commun, vol. 5, p. 3071, 2014. View at: Google Scholar
                          155. L. Arnold, A. Henry, F. Poron et al., “Inflammatory monocytes recruited after skeletal muscle injury switch into antiinflammatory macrophages to support myogenesis,” The Journal of Experimental Medicine, vol. 204, no. 5, pp. 1057–1069, 2007. View at: Publisher Site | Google Scholar
                          156. J. E. Heredia, L. Mukundan, F. M. Chen et al., “Type 2 innate signals stimulate fibro/adipogenic progenitors to facilitate muscle regeneration,” Cell, vol. 153, no. 2, pp. 376–388, 2013. View at: Publisher Site | Google Scholar
                          157. D. Burzyn, W. Kuswanto, D. Kolodin et al., “A special population of regulatory T cells potentiates muscle repair,” Cell, vol. 155, pp. 1282–1295, 2013. View at: Publisher Site | Google Scholar


                          Copyright © 2018 Juan Liu et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

                          Can Titanium Plates Be Left in the Body Safely?

                          Patients who have metal plates, pins, and screws in the body are rightfully concerned about the safety and long-term effects of these devices left inside. Surgeons share those concerns but do not want to perform an additional surgery to remove them if unnecessary. And sometimes when plates are used to hold bone together after a fracture, bone grows around them embedding this fixation device too much to remove it easily.

                          So just how s
                          afe are these devices? In this article, Dr. David G. Dennison from the Orthopedic Surgery department of Mayo Clinic in Rochester, Minnesota summarizes what we know from research and clinical studies on this topic. In particular, Dr. Dennison zeroes in on titanium volar plates used to treat distal radius fractures.

                          Titanium has replaced stainless steel these days for fixation devices. It is more compatible with the human biology, which means it’s less likely to cause a reaction. When an inflammatory response does occur, it is mild and limited (doesn’t last). Titanium can also be combined with other metals such as cobalt, chromium, and molybdenum to create a lighter but more durable material.

                          Volar plates refer to the location of fixation devices — placed on the front or inside of the forearm. Radius fracture tells us the radial bone in the forearm is broken. There are two bones in the forearm: ulna and radius. The radius is on the thumb-side of the forearm. Distal means the break is down toward the hand rather than up by the elbow.

                          There are all kinds of concerns about metal plates. Animal studies show there is an effect on the immune system. There is evidence that the metal can cause the entire immune system to be suppressed (under functioning). This immune system shut down could result in infections. Some studies have shown that metal implants can cause an increase in white blood cells called lymphocyte reactivity. There is a worry that this effect could cause implant loosening or failure, though it hasn’t been proven yet.

                          Another potential problem with titanium plates is the debris that occurs. Tiny flakes of this metal chip off and enter the bloodstream, nearby soft tissues, and/or joint. Both titanium and stainless steel have been found in all these anatomical areas of the human body (titanium slightly more often and in greater amounts than stainless steel). Metal debris is more likely to develop when the implant is rubbing against another surface. This wearing or rubbing phenomenon is called fretting.

                          Then the question arises: can this metal debris lead to the formation of cancer? Studies in mice show there is the potential for metal wear debris to damage chromosomes making it a potential carcinogen (cancer producing). Next, developers of these products asked if coating the plate would protect the body from corrosion or metal debris? This question remains unanswered so far.

                          One thing we do know from studies — placing a long titanium or metal pin down through the middle of a bone to stabilize it is linked with a much higher increase in the amount of metal found in the bloodstream. Chromium seems to have the highest levels reported for these intramedullary nails. Intramedullary titanium nails also increase the amount of titanium found in blood samples, but not as much as chromium. Evidently, the large surface area of the intramedullary nail exposes the bone to more titanium, thus the higher levels of serum (blood) metal.

                          Removing titanium plates does slowly reduce the levels of metal in the blood and soft tissues. But it can take a year or more after removal to bring these elevated levels back down to normal. Dr. Dennison reports that there aren’t specific studies of the effects of titanium plates (prolonged use or removal) from the treatment of distal volar radial fractures.

                          Summarizing, Dr. Dennison says that the information found on this topic seems to suggest that the overall level of risk when leaving titanium plates in the forearm is acceptable. The volar surface plate is not a large implant and doesn’t usually involve nails down through the bone. Instead, small screws placed perpendicular to the plate to hold it in place are more typical. Still, there are concerns and risks reported that deserve specific study and more conclusive answers. Future studies are needed to address these concerns and questions.

                          Why wounds heal more slowly with age

                          Older bodies need longer to mend. This reality of aging has been documented since World War I, with the observation that wounds heal slower in older soldiers. Yet until now, researchers have not been able to tease out what age-related changes hinder the body's ability to repair itself.

                          Recent experiments at The Rockefeller University explored this physiological puzzle by examining molecular changes in aging mouse skin. The results, described November 17 in Cell, delineate a new aspect of how the body heals wounds.

                          "Within days of an injury, skin cells migrate in and close the wound, a process that requires coordination with nearby immune cells. Our experiments have shown that, with aging, disruptions to communication between skin cells and their immune cells slow down this step," says Elaine Fuchs, the Rebecca C. Lancefield Professor and head of the Robin Chemers Neustein Laboratory of Mammalian Cell Biology.

                          "This discovery suggests new approaches to developing treatments that could speed healing among older people," adds Fuchs, who is also a Howard Hughes Medical Institute investigator.

                          Return of the skin cells

                          Whenever a wound occurs, the body needs to repair it quickly to restore its protective skin barrier. "Wound healing is one of the most complex processes to occur in the human body," says Brice Keyes, a former postdoc in Fuch's lab and currently a researcher at Calico Life Sciences. "Numerous types of cells, molecular pathways, and signaling systems go to work over timescales that vary from seconds to months. Changes related to aging have been observed in every step of this process." Keyes and Siqi Liu, an immunology specialist and a current Jane Coffin Childs postdoctoral fellow in in the lab, are co-first authors of the Cell article.

                          Both skin cells and immune cells contribute to this elaborate process, which begins with the formation of a scab. New skin cells known as keratinocytes later travel in as a sheet to fill in the wound under the scab.

                          The team focused on this latter step in healing in two-month-old versus 24-month-old mice -- roughly equivalent to 20- and 70-year-old humans. They found that among the older mice, keratinocytes were much slower to migrate into the skin gap under the scab, and, as a result, wounds often took days longer to close.

                          Wound healing is known to require specialized immune cells that reside in the skin. The researchers' new experiments showed that following an injury, the keratinocytes at the wound edge talk to these immune cells by producing proteins known as Skints that appear to tell the immune cells to stay around and assist in filling the gap. In older mice, the keratinocytes failed to produce these immune signals.

                          Seeking a reversal

                          To see if they could enhance Skint signaling in older skin, the researchers turned to a protein that resident immune cells normally release after injury. When they applied this protein to young and old mouse skin tissue in a petri dish, they saw an increase in keratinocyte migration, which was most pronounced in the older skin. In effect, the old keratinocytes behaved more youthfully.

                          The scientists hope the same principle could be applied to developing treatments for age-related delays in healing.

                          "Our work suggests it may be possible to develop drugs to activate pathways that help aging skin cells to communicate better with their immune cell neighbors, and so boost the signals that normally decline with age," Fuchs says.


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