Simple Marinara Sauce | My Bizzy Kitchen

By electricdiet / August 28, 2021

This simple marinara sauce takes only 20 minutes to make, yet is rich and tastes like it’s been simmering all day.  Tomato paste gives this a depth of flavor that is delicious.

this is a photo of simple marinara sauce

A few months ago I bought a Vitamix on sale for $249.  I am kicking myself for not buying one sooner – it’s amazing.  Costco sometimes has Vitamix’s on sale too.

this is a photo of a Vitamix blending simple marinara sauce

What is the secret to a simple marinara?

Good canned tomatoes.  My late husband introduced me to peeled Italian tomatoes and they are so delicious and readily available in any grocery store.  The cans cost between $1.49 and $2.00 not on sale, but about every three weeks they are on sale for $.99 so I stock up.

What can you do with leftover marinara sauce?

I live alone, but I still make this full batch to use throughout the week.  Here are some recipes that use marinara sauce:

You can use some of this marinara sauce to make my chicken meatball dish.

this is a photo of chicken meatballs with marinara

Zucchini Lasagna with Pork Ragu:

this is a photo of zucchini lasagna

Quick Tomato Soup:

this is a photo of quick tomato soup


Or my quick chicken parmesan.  

4 ounces chicken breast tossed in 1 tablespoon of flour, salt and pepper.  Pan fried over medium heat for 4 minutes a side.  On the last minute, top with 1 tablespoon shredded Parmesan cheese and put that side down in the pan so it gets nice and crispy.  Serve over whole wheat pasta and some of the marinara.  Garnish with fresh basil.

This is a photo of crispy chicken parmesan over whole wheat pasta


  • 1 28 ounce can Italian peeled plum tomatoes
  • 2 tablespoons minced garlic
  • 2 tablespoons tomato paste
  • 1 tablespoon oregano
  • 1 tablespoon Italian seasoning
  • 1/2 teaspoon crushed red pepper
  • 1/2 teaspoon salt
  • 1/2 teaspoon pepper


  1. Spray a skillet with avocado oil spray. Add the garlic and cook for five minutes. Add the tomato paste and cook for two minutes. Add the remaining ingredients and simmer for 10 minutes.
  2. Use a blender or stick blender, puree sauce.


This is ZERO points on all WW plans. If your sauce is a bit acidic, add a pinch of sugar to balance it out. A teaspoon of sugar is only 10 calories, so I wouldn’t count that. 😂

Nutrition Information:

Yield: 4

Serving Size: 1

Amount Per Serving:

Calories: 55Total Fat: 1gSaturated Fat: 0gTrans Fat: 0gUnsaturated Fat: 0gCholesterol: 0mgSodium: 281mgCarbohydrates: 12gFiber: 4gSugar: 6gProtein: 3g

Did you make this recipe?

Please leave a comment on the blog or share a photo on Instagram

Let me know if you make this – I’d love to hear what you think!

This homemade marinara sauce takes only 20 minutes to make, yet it is rich and tastes like it’s been simmering all day. Tomato paste gives this a depth of flavor that is delicious. You can use some of this marinara sauce to make my chicken meatball recipe, my zucchini lasagna, my quick tomato soup, and my chicken parmesan. This is ZERO points on all WW plans. If your sauce is a bit acidic, add a pinch of sugar to balance it out. #ww #weightwatchers #pasta #marinara #sauce


I am a widowed 52 year old trying to figure out my life after losing my husband after a long illness.

Cooking and being in the kitchen feeds my soul!

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Basaglar vs Lantus: What’s The Difference?

By electricdiet / August 26, 2021

While there are many different types of insulin glargine, two of the most popular are Lantus and Basaglar. 

Lantus was introduced to the market in 2000 and Basaglar was introduced as a “biosimilar” option in 2015 

Lantus and Basaglar are both prescribed to treat patients with type 1 and type 2 diabetes and are very similar, but not always interchangeable.

This article will explain these two popular types of insulin, their uses, and which may be a better option for you. 

The need for basal insulin 

All people, whether or not they have diabetes, require insulin to live. People with diabetes, however, either cannot make their own insulin, or their body fails to use the insulin their body produces properly (some people with type 2 experience this in the form of insulin resistance). 

People who choose not to use insulin pump therapy and are thus on multiple daily injections (MDI) will require both basal and bolus insulins. 

Basal insulin or “background” insulin has one primary job: to keep your blood sugars level when you’re not eating (when you’re fasting) and overnight when you’re asleep. 

Long-acting (basal) insulin is unique in the sense that it slowly releases the insulin you inject over a longer period of time, as opposed to short-acting insulin which releases and is absorbed much faster in the body.

Long-acting insulin works well with bolus insulins (short-acting insulin) that is taken for all foods (especially carbohydrates) consumed.

Since all humans require insulin all the time to live, even without eating anything, a person with diabetes will experience severe hyperglycemia (high blood sugar) if they do not take basal insulin. 

There are different categories of long-acting insulin:

  1. Intermediate-acting insulin (NPH), peaks at 8 hours, lasts 12-16 hours 
  2. Long-acting insulin (detemir or glargine), no peak, lasts about 24 hours 
  3. Ultra-long acting insulin (degludec), no peak, lasts up to 42 hours 

The most common form of long-acting insulin is glargine. There are two brand names of glargine available: Basaglar and Lantus. 

What is Lantus? 

Lantus was first developed by the insulin manufacturer Sanofi in 2000 and is used to treat both type 1 and type 2 diabetes. 

Lantus was one of the first longer-acting insulins on the market, making modern-day carbohydrate counting, taking insulin for food eaten (instead of the other way around), and insulin to carbohydrate ratios a reality. 

The introduction of Lantus was revolutionary to people on multiple daily injections, as adding in a basal insulin got rid of the necessary snacking throughout the day that was required to counter the peaks of intermediate-acting insulins like NPH. 

The introduction of Lantus on the market greatly decreased the number of nocturnal hypoglycemic events people with diabetes experienced, due to its lack of a peak, and its steady, slow release of insulin over a 24 hour period, and improved the lives of many people living with diabetes. It is still a very popular option for people who are on MDI. 

Lantus comes in both an insulin vial and insulin pen version. 

What is Basaglar? 

Basaglar, developed by Eli Lilly & Company, was first introduced in 2015. The reason for the 15-year delay was that Lantus was patent-protected for the first 15 years, and until those patents expired, other insulin manufacturers could not make similar insulins that would compete for market share. 

Basaglar was the first “biosimilar” insulin to be approved by the Food and Drug Administration as well as the first to launch in the United States, paving the way for more affordable options on the insulin market. 

Other pharmaceutical companies, such as Mylan/Biocon and Merk/Samsung Bioepis are also working on biosimilar glargine insulins as well, which should drive down prices as competition increases. 

Similar to Lantus, Basaglar has a slow release over a 24 hour period and has no peak. 

While the two insulins perform remarkably similarly (including the same protein sequence and blood sugar-lowering effects), their chemical formularies are ever so slightly different, so people with diabetes cannot simply switch out one type of insulin for the other without their doctor’s guidance. 

Basaglar is only available via an insulin pen called the Kwikpen. 

What is the difference between these two insulins? 

While these insulins are remarkably similar, people may react differently to their chemical makeup. 

Adverse events such as slower absorption times, skin irritation, or seeing a slight peak (although uncommon, it may happen) may happen to you if you try one insulin and not if you try the other. 

Always work with your provider to see what the best option is for you. 

Another main difference between these two insulins is the cost. The out-of-pocket cost for each in the United States is:

  • Lantus: $425 for 5 pens (300 units a piece) or around $285 for one vial (1,000 units)
  • Basaglar: $326 for 5 pens (300 units a piece) 

This makes sense, as Basaglar was introduced as a more affordable option to Lantus. 

Is Lantus or Basaglar more effective for blood sugar control?

The two insulins are equally effective for blood sugar control. Any differences you may experience are the result of your individual reaction to the insulin and not because one brand is superior to the other. 

Can you substitute Lantus for Basaglar? 

Basaglar is not a generic equivalent of Lantus, and therefore, the two are not interchangeable. 

Although Lantus and Basaglar contain the same active ingredient, they are made differently and by different companies. This means that Basaglar is not an exact replica of Lantus. 

If you are prescribed one and would like to switch to the other, your doctor will need to write you a new prescription. Your pharmacist cannot just substitute one for the other without your doctor’s approval.

What are the side effects of these two insulins? 

The main side effect of these two insulins is hypoglycemia (low blood sugar). Symptoms of hypoglycemia include:

  • Headache
  • Weakness
  • Sweating
  • Increased heart rate 
  • Troubling concentrating 
  • Hunger 
  • Fainting
  • Shakiness 
  • If severe, seizure 

Other common side effects of glargine include: 

  • Skin irritation at the injection site
  • Redness at the injection site 
  • Pain and swelling at the injection site 
  • Thickening or hollowing of the skin at the injection site 

Speak with your doctor if you’re pregnant, planning on becoming pregnant or breastfeeding and wish to start glargine. Your insulin requirements will vary during and after pregnancy. 

Which one is the right type of insulin for me? 

There are many reasons why people choose one of these insulins over the other. You may have a very specific preference for the type of long-acting insulin that you choose. 

Perhaps you do not want to use an insulin pen (and you’ll want to go with Lantus), or perhaps you will ONLY use an insulin pen (making Basaglar your insulin of choice). 

Maybe you experience an adverse reaction to one type of insulin and not the other (pain, swelling, itching, redness or swelling at the injection site are common side effects of both Lantus and Basaglar). 

Perhaps your body metabolizes one insulin better than the other. Some people may find that one insulin or the other doesn’t quite last a full 24 hours, and they need to split their dose

To know for sure, you should work with your provider to see what they recommend. 

Talk about your lifestyle and health goals, and seek their advice, but both insulins come highly rated, and choosing the best one for you and your diabetes management is ultimately up to you! 

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Diabetic Friendly Desserts – Berry Parfait Tops My Best Berry Desserts

By electricdiet / August 24, 2021

Yes It’s True – Delicious Diabetic Friendly Desserts!

Diabetic friendly desserts are amazing and especially when berries are in season. We have the best berry desserts and couldn’t wait to share them. Holly Clegg’s super easy diabetic recipes like this Berry Parfait from her arthritis cookbook will wow you! Eating Well To Fight Arthritis cookbook includes healthy easy recipes to help manage an anti-inflammatory diet.  Berries are high in antioxidants making them good arthritis diet recipes.  You should enjoy food and make healthy eating with nutrition a lifestyle. You’ll love this luscious berry dessert and how quick you can make it.

berry parfait diabetic

Eating Diabetic Friendly Desserts

A diabetic diet is the healthiest way to eat and not a diet! These easy diabetic recipes include your favorite foods. Best of all, these desserts and diabetic recipes don’t taste like your sacrificing flavor.  This berry dessert makes the perfect ending to any meal. Team Holly’s cookbooks include mainstream easy diabetic recipes!

Fresh Juicy Berries Make Best Berry Desserts

Berries can be found year round.  However, when strawberries are in season, put this fantastic sweet treat high on your list. We love using berries in desserts. This Berry Parfait is fabulous but you can use your favorite seasonal berry combination also. Garnish with an impressively easy curl of orange peel. You can serve this gorgeous parfait in individual dishes for extra pizzazz and presentation.  Like these inexpensive glass bowls.  Here’s a fun gadget to hull strawberries and its an inexpensive strawberry huller. 

EATING WELL TO FIGHT ARTHRITISEATING WELL TO FIGHT ARTHRITISEATING WELL TO FIGHT ARTHRITISGodinger Dublin Set Of 4 Candy BowlsGodinger Dublin Set Of 4 Candy BowlsGodinger Dublin Set Of 4 Candy BowlsJoie Stainless Steel Strawberry HullerJoie Stainless Steel Strawberry HullerJoie Stainless Steel Strawberry Huller

Berry Parfait
Light and delightful! Use strawberries or your favorite berries. Then, layers a luscious cream cheese filling and lady fingers or angel food cake and makes one of the best berry desserts. Also, this easy berry recipe makes a great make ahead dessert. I especially love it for summer recipes but I make it all year long.

    Servings16 (1/2 cup) servings


    • 2/3cup

      spreadable seedless sugar free raspberry jam

    • 1/4cup

      orange liqueur or orange juicedepending on taste

    • 6ounces

      reduced-fat cream cheese

    • 1/4cup


    • 1teaspoon

      vanilla extract

    • 1teaspoon

      grated orange rind

    • 1(8-ounce) container

      free fat free whipped toppingthawed

    • 2(3-ounce) packages

      ladyfingerssplit in half (24) or angel food cake cut into pieces

    • 1/2cup


    • 1 1/2cups

      strawberriesstemmed and sliced

    1. In small bowl, mix together preserves and orange juice.

    2. In mixing bowl, cream together cream cheese, sugar, and vanilla until light.  Mix in whipped topping and reserve 1/2 cup for topping.

    3. Line bottom of an oblong small dish with split ladyfingers. Layer with preserves mixture, cream cheese layer, and top with strawberries. Repeat layers with remaining ladyfingers, preserves and cream cheese layer.

    4. Carefully top with thin layer of remaining whipped topping. Top with remaining strawberries. Cover, refrigerate several hours or overnight.

    Recipe Notes

    Calories 128, Calories from fat (%) 24, Fat (g) 3, Saturated Fat (g) 1, Cholesterol (mg) 9, Sodium (mg) 79, Carbohydrate (g) 23, Dietary Fiber (g) 1, Sugars (g) 11, Protein (g) 2, Diabetic Exchanges: 1 1/2 carbohydrate, 1/2 fat

    berry parfait diabetic

    Get Cookbook With Cooking Tips To Simplify Your Kitchen

    Terrific Tip: Substitute raspberries or blueberries for strawberries.  Then,  if you are having joint pain you won’t have to slice strawberries. Try using a berry mixture. Lower the recipe’s sugar content by using sugar-free angel food cake. If desired, leave out the orange rind or find dried orange rind in the spice section.

    Eating Well Through Arthritis is filled with tips to simplify your cooking and recipes!  You’ll love the healthy easy recipes and you probably already have the ingredients!  Not only does this cookbook contain arthritis diet recipes but it is truly everyday recipes. Start healthy cooking today with familiar favorite recipes.


    Peanut Butter Cookies! Popular easy diabetic recipe and all time favorite peanut butter cookie recipe!

    What to do with all those berries?! Find more berry recipes on Team Holly’s healthy food blog. Ever had Berry Good Oatmeal Cookie Cake? OMG!  Red, White and Blue recipes  make fun patriotic desserts.  Celebrate patriotic holidays with berries for healthy eating.  Besides, berries not only make easy diabetic desserts but they make the best berry dessert. See you in the kitchen.

    Get All of Holly’s Healthy Easy Cookbooks

    The post Diabetic Friendly Desserts – Berry Parfait Tops My Best Berry Desserts appeared first on The Healthy Cooking Blog.

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    Glucagon-Like Peptide 1 Secretion by the L-Cell

    By electricdiet / August 22, 2021

    The View From Within


    Glucagon-like peptide 1 (GLP-1) is a gut-derived peptide secreted from intestinal L-cells after a meal. GLP-1 has numerous physiological actions, including potentiation of glucose-stimulated insulin secretion, enhancement of β-cell growth and survival, and inhibition of glucagon release, gastric emptying, and food intake. These antidiabetic effects of GLP-1 have led to intense interest in the use of this peptide for the treatment of patients with type 2 diabetes. Oral nutrients such as glucose and fat are potent physiological regulators of GLP-1 secretion, but non-nutrient stimulators of GLP-1 release have also been identified, including the neuromodulators acetylcholine and gastrin-releasing peptide. Peripheral hormones that participate in energy homeostasis, such as leptin, have also been implicated in the regulation of GLP-1 release. Recent studies have begun to elucidate the intracellular signaling pathways that mediate the effects of GLP-1 secretagogues on the intestinal L-cell. The purpose of this review is to summarize the known signaling mechanisms of GLP-1 secretagogues based on the available literature. A better understanding of the pathways underlying GLP-1 secretion may lead to novel approaches by which the levels of this important insulinotropic hormone can be enhanced in patients with type 2 diabetes.

    Glucagon-like peptide 1 (GLP-1) is an intestinal hormone that exerts profound effects in the regulation of glycemia, stimulating glucose-dependent insulin secretion, proinsulin gene expression, and β-cell proliferative and anti-apoptotic pathways, as well as inhibiting glucagon release, gastric emptying, and food intake (1). The demonstrated success of GLP-1 to lower glycemia has led to approval of the GLP-1 receptor agonist exendin-4 (Byetta) for the treatment of patients with type 2 diabetes (2). Studies using GLP-1 receptor antagonists as well as GLP-1 receptor null mice have demonstrated that GLP-1 makes an essential contribution to the “incretin” effect after a meal (3,4). However, GLP-1 secretion is reduced in patients with type 2 diabetes (57), and this may contribute in part to the reduced incretin effect and the hyperglycemia that is observed in these individuals (8). Thus, interest has now focused on the factors that regulate the release of this peptide after nutrient ingestion. Many different GLP-1 secretagogues have been described in the literature over the past few decades, including nutrients, neurotransmitters, neuropeptides, and peripheral hormones (rev. in 9,10). However, the specific receptors, ion channels, and intracellular signaling proteins expressed by the GLP-1–producing intestinal L-cell have only recently begun to be characterized. The purpose of this review is to integrate the literature regarding the signal transduction pathways used by the major GLP-1 secretagogues. An improved understanding of the mechanisms regulating GLP-1 secretion may lead to novel approaches to enhance GLP-1 levels in vivo, thereby providing an alternative approach to the use of this peptide in the treatment of patients with type 2 diabetes.


    Although the proglucagon gene is expressed in enteroendocrine L-cells and pancreatic α-cells (11), GLP-1 is synthesized by posttranslational processing of proglucagon only in the intestine. The L-cells are predominantly located in the ileum and colon and have been identified as open-type epithelial cells that are in direct contact with nutrients in the intestinal lumen (12). Furthermore, L-cells are located in close proximity to both neurons and the microvasculature of the intestine (13,14), which allows the L-cell to be affected by both neural and hormonal signals. In fetal rat intestinal L-cell cultures and immortalized murine L-cells, proglucagon gene expression is enhanced by activation of the protein kinase A (PKA) pathway (15,16). Although originally thought to be mediated through CREB binding to the cAMP response element in the proglucagon gene, recent studies have demonstrated that cAMP-dependent proglucagon gene expression in the L-cell occurs via β-catenin–mediated activation of the bipartite transcription factor, TCF4 (17). Interestingly, a very recent report has linked variants of TCF4 to the risk for development of type 2 diabetes (18), implicating GLP-1 in the etiology of this disease.

    Tissue-specific expression of prohormone convertase isoforms directs the synthesis of specific proglucagon-derived peptides in the L-cell and α-cell (Fig. 1). Hence, cleavage of proglucagon by prohormone convertase 1/3, which is expressed in the L-cell, liberates GLP-1 and GLP-2, as well as the glucagon-containing peptides, glicentin and oxyntomodulin (19,20). In contrast, α-cell expression of prohormone convertase 2 leads to synthesis of glucagon, glicentin-related pancreatic peptide, and the major proglucagon fragment, which contains within its sequence both GLP-1 and GLP-2. Early studies on L-cell secretagogues used antibodies that targeted a mid-sequence epitope of glucagon, resulting in detection of glicentin/oxyntomodulin (i.e., enteroglucagon or gut glucagon-like immunoreactivity) as well as of glucagon. Because these peptides, as well as GLP-2, are synthesized and secreted from the L-cell in a 1:1 stoichiometric ratio with GLP-1 (21,22), such studies therefore indirectly also examined GLP-1 synthesis and secretion. Hence, the results of studies using antisera against any of the intestinal proglucagon-derived peptides are discussed in this review synonymously. Finally, bioactive GLP-1 exists in two equipotent forms, GLP-17-36NH2 and GLP-17-37, in the circulation, of which the former is predominant (23). Secreted GLP-1 is rapidly degraded by the ubiquitous enzyme dipeptidyl peptidase IV (24), resulting in an extremely short half-life for GLP-1 of ∼2 min (23).


    Nutrient ingestion is the primary physiological stimulus to the L-cell and results in a biphasic pattern of GLP-1 secretion. An initial rapid rise in circulating GLP-1 levels occurs 15–30 min after a meal, followed by a second minor peak at 90–120 min (25). Glucose and fat have been found to be potent stimulators of GLP-1 secretion when ingested (26), but also after direct administration into the intestinal lumen (22,27) or into perfused ileal segments (28). Unlike glucose and fat, protein does not appear to stimulate proglucagon-derived peptide secretion from L-cells (26), although protein hydrolysates have been found to stimulate GLP-1 release in a perfused rat ileum model and in immortalized human L-cells (28,29).

    Previous studies have indicated that L-cells are not present in the proximal small intestine of humans and rodents (12). Hence, we proposed that the initial rapid rise in GLP-1 secretion must be mediated indirectly, through a neuro/endocrine pathway, rather than through direct interactions of the luminal contents with L-cells (27). Although a very recent study has detected GLP-1–immunoreactive cells in the duodenum of humans (30), this remains controversial. Therefore, to elucidate the mechanism(s) underlying proximal nutrient-stimulated GLP-1 secretion, we developed a rodent model of GLP-1 secretion in which nutrient flow to the distal intestine was prevented, thereby excluding the possibility of direct interactions of the luminal nutrients with the L-cell (Fig. 2). In this model, placement of glucose or fat into the duodenum induced an immediate and prolonged stimulation of the L-cell that was comparable in magnitude to increments in proglucagon-derived peptides observed when nutrients were placed directly into the ileum (27). Furthermore, when nutrients were placed in the duodenum of the rat, a prompt rise in glucose-dependent insulinotropic peptide (GIP) levels was also observed, and infusion of GIP or treatment of primary rat L-cells in culture with GIP also stimulated proglucagon-derived peptide secretion (22,27,31), thus implicating GIP in the proximal regulation of GLP-1 secretion. The more important role of the vagus nerve in mediating the proximal-distal loop was elucidated when L-cell stimulation by placement of fat into the duodenum or by infusion of physiological concentrations of GIP was completely abrogated by subdiaphragmatic vagotomy (22). Furthermore, activation of the efferent celiac branch was demonstrated to increase GLP-1 secretion. Finally, acetylcholine has now been identified as a key neurotransmitter mediating the proximal-distal loop. In vitro studies have determined that human and rodent L-cells are sensitive to the stimulatory effects of acetylcholine (13,32). Furthermore, when rodents were infused with atropine or pirenzipine (an M1 muscarinic receptor antagonist), duodenal nutrients were unable to increase GLP-1 secretion (13). When taken together, therefore, these studies demonstrated that, in rodents, the regulation of GLP-1 release by proximal nutrients is mediated via GIP actions on cholinergic fibers of the vagus nerve.

    A second neural regulator of the proximal-distal loop has been identified as gastrin-releasing peptide (GRP), a neuropeptide that is locally released from GRPergic neurons in the enteric nervous system. GRP has been demonstrated to be a potent GLP-1 secretagogue in both in vitro and in vivo studies of the L-cell (31,33). The role of GRP in the proximal-distal loop regulating the L-cell has also been demonstrated using a GRP receptor antagonist, as well as GRP null mice (33,34).

    Although the early rise of GLP-1 does occur in humans (25,26), the specific mediators of the proximal-distal loop have yet to be defined. The human L-cell is responsive to acetylcholine in vitro (32), and administration of atropine prevents GLP-1 secretion after nutrient ingestion (35). Reimer et al. (29) have also demonstrated that human L-cells are responsive to GRP in vitro. However, GIP does not stimulate GLP-1 secretion in humans in vivo (36). Further work is clearly required to identify the exact mechanisms mediating the rapid release of GLP-1 secretion after nutrient ingestion in humans.

    The second later peak of GLP-1 secretion is believed to occur consequent to transit of ingested nutrients down the lumen to directly interact with the distal L-cell. Placement of nutrients directly into the lumen of the ileum stimulates GLP-1 release (27,37), whereas treatment of rodent and human L-cells in culture with either glucose or fatty acids induces dose-dependent increases in GLP-1 secretion (29,3841). As glucose does not reach the distal gut in high concentrations, it has been proposed that fat, which does transit to the ileum, is the more physiological direct regulator of GLP-1 release (9). Nonetheless, under conditions of nutrient dumping/rapid transit or after administration of digestive enzyme inhibitors, both fat- and glucose-stimulated GLP-1 secretion may be of importance (4244).

    Finally, in addition to luminal nutrients, a number of additional intestinal peptides and neurotransmitters, as well as systemic hormones, have been demonstrated to modulate GLP-1 secretion. These include intestinal somatostatin, for which a negative feedback loop with GLP-1 has been proposed (31,45); the neurotransmitter γ-aminobutyric acid (GABA), which enhances GLP-1 release (46); and α-and β-adrenergic agonists, which respectively inhibit and/or stimulate GLP-1 secretion from L-cells in the perfused rat intestine (47). More recently, the adipocyte hormone leptin has also been found to stimulate GLP-1 release from the human and rodent L-cell, and this effect is abolished in leptin-resistant diet-induced obese mice (48). Furthermore, preliminary findings have demonstrated insulin to be a GLP-1 secretagogue, while insulin resistance in the L-cell reduces basal and stimulated GLP-1 release (49). Although the physiological significance of many of these findings remains to be firmly established, these studies suggest that impairments at the level of the L-cell may account, at least in part, for the reduced GLP-1 secretion that is observed in patients with type 2 diabetes (57), as well as in obesity (50).


    Until recently, there has been a relative paucity of published literature on the intracellular pathways mediating the effects of GLP-1 secretagogues in the L-cell. However, the development of in vitro models of the murine, rat, and human intestinal L-cell has now permitted more detailed examination of signaling pathways expressed in the L-cell, although in some instances, information has been assumed by extrapolation from what is known regarding other neuroendocrine cells (Figs. 35). An improved understanding of the intracellular mechanisms underlying GLP-1 secretion may lead to novel approaches to augment GLP-1 secretion in vivo, particularly if used in conjunction with agents designed to prevent the rapid degradation of GLP-1 in the circulation (1).


    As discussed above, acetylcholine stimulates GLP-1 secretion in vitro and in vivo. M1 muscarinic receptor agonists increase GLP-1 secretion from rat L-cells, and inhibition of M1 muscarinic receptors prevents proximal nutrient-induced release of GLP-1 by bethanechol, a nonselective muscarinic agonist (13). In human L-cells, inhibition of M1 and M2 muscarinic receptors also prevents bethanechol-induced GLP-1 release (32). M1 muscarinic receptors are G protein–coupled receptors that are linked to Gαq/11, and ligand binding results in the activation of phospholipase C, which cleaves phosphatidylinositol (4,5)biphosphate into inositol-1,4,5-trisphosphate and diacylglycerol, leading to increases in intracellular calcium and activation of both conventional and novel protein kinase C (PKC) isoforms (51) (Fig. 3). M2 receptors are thought to be coupled to Gαi, which inhibits adenylyl cyclase (51), but the enhanced GLP-1 secretory response to M2 receptor activation in human L-cells suggests the existence of an alternative intracellular pathway.


    GRP is a potent stimulator of the intestinal L-cell in vivo and in vitro (31,33), but the signal transduction cascade that occurs in response to GRP treatment in the L-cell has yet to be defined. Based on studies using other neuroendocrine cells, GRP binds to a G protein–coupled receptor that is coupled to Gαq (52). For example, the plurihormonal murine secretin tumor cell line (STC-1) releases not only secretin, but also GLP-1 and cholecystokinin. Treatment with GRP stimulates hormone secretion by these cells in association with activation of mitogen-activated protein kinase kinase (MAPKK) and subsequent phosphorylation of p44/42 mitogen-activated protein kinase (MAPK). GRP-stimulated cholecystokinin secretion was also found to be dependent on the activation of PKC (53). Consistent with these findings, downregulation of PKC activity by prolonged treatment with phorbol myristate acetate to inactivate classic and novel PKCs prevents GRP-mediated insulin secretion from pancreatic β-cells (54). GRP also enhanced insulin secretion in association with an increase in intracellular calcium. Although p44/42 MAPK is expressed in the mouse and human L-cell (49; R. Iakoubov, A. Izzo, A. Yeung, C.I. Whiteside, P.L.B., unpublished data) and changes in intracellular calcium levels have been linked to GLP-1 release in the rodent L-cell (55,56), further work is clearly required to determine the exact mechanism of action of GRP to stimulate GLP-1 secretion.


    GABAergic neurons are components of the enteric nervous system located primarily in the myenteric plexus of the colon. Three isoforms of the GABA receptor exist (GABAA, GABAB, and GABAC), and their expression and distribution is tissue specific. Of the three isoforms, GABAA and GABAC receptors are ion-channel linked receptors, whereas the GABAB receptor is a metabotropic G protein–coupled receptor (57). Gameiro et al. (46) confirmed the expression of GABAA receptors in the murine L-cell, and GABA treatment of these cells caused an efflux of chloride ions from the cell, leading to depolarization, opening of voltage-gated calcium channels, and GLP-1 secretion. These in vitro findings suggest that GABA from GABAergic neurons may act in a paracrine manner to modulate hormone secretion. Nonetheless, the physiological role of GABA modulation of GLP-1 secretion in vivo still remains to be demonstrated.

    Glucose-dependent insulinotropic peptide.

    GIP mediates its biologic actions through a G protein–coupled receptor belonging to the glucagon receptor superfamily, which includes receptors for other structurally related gut-derived peptides, including GLP-1, GLP-2, glucagon, secretin, and growth hormone–releasing hormone (58). GIP receptor activation in the β-cell leads to the activation of adenylyl cyclase through Gαs, resulting in increases in cAMP as well as in cytosolic calcium (59). This pathway leads to downstream activation of PKA and enhances hormone release, most notably that of insulin from the β-cell (60). However, GIP has also been reported to stimulate insulin secretion through cAMP-dependent PKA-independent activation of the cAMP guanine nucleotide exchange factor-II (Epac2) pathway (61). Although a direct stimulatory effect of GIP on the canine and rodent L-cell has been demonstrated (31,39,62), the expression of the GIP receptor and its downstream signaling pathways has yet to be fully examined. Furthermore, although cAMP-dependent GLP-1 secretion has been demonstrated in the human, murine, and rat L-cell (29,39,56), whether this is modulated through a PKA-dependent pathway and/or Epac2 has yet to be determined.


    Two distinct forms of somatostatin are produced in the intestine, SS14 by enteric neurons, and SS28 by enteroendocrine D-cells. However, in both rats and pigs, SS-28 is a more potent inhibitor of GLP-1 secretion (31,63,64). Somatostatin receptors exist as five isoforms (sst1–sst5), of which sst5 is expressed by the rat L-cell (63). These receptors are coupled with a pertussis-sensitive Gαi protein, and activation both inhibits adenylyl cyclase and decreases intracellular calcium levels (65). Because GLP-1 stimulates the secretion of both forms of somatostatin from the intestine, these findings suggest the existence of a feedback loop through which locally produced intestinal somatostatin can modulate GLP-1 release after the ingestion of nutrients (31,45,63,64).

    Fatty acids.

    Long-chain monounsaturated fatty acids (MUFAs) directly stimulate GLP-1 secretion from the murine, rat, and human L-cell (29,38,39). Several recent studies have now begun to elucidate the mechanism of action of fatty acids on the L-cell (Fig. 4). Recently, long-chain fatty acids have been found to interact with two distinct orphan G protein–coupled receptors: GPR40 and GPR120. In the β-cell, GPR40 is coupled to both Gαq and Gαi, as demonstrated by increases in cytosolic calcium and inhibition of forskolin-induced cAMP production, respectively (66). GPR40 activation also increases p44/42 MAPK phosphorylation, and siRNA-mediated knock-down of GPR40 prevents fatty acid–induced insulin secretion. mRNAs for both GPR40 and GPR120 have also been detected in the murine L-cell, as well as in STC-1 cells (R. Iakoubov, A. Izzo, A. Yeung, C.I. Whiteside, P.L.B., unpublished data; 67). Although GPR120, but not GRP40, is required for fatty acid–induced GLP-1 release from the STC-1 cells, and activation of GPR120 is linked to increases in cytosolic calcium as well as p44/42 MAPK phosphorylation in these cells, neither of these pathways appear to be required for fatty acid–induced GLP-1 release (67). Furthermore, despite expression of both GPRs in the murine L-cell, MUFA treatment of these cells does not increase either intracellular calcium levels or p44/42 MAPK phosphorylation (R. Iakoubov, A. Izzo, A. Yeung, C.I. Whiteside, P.L.B., unpublished data). Hence, the exact role of these novel fatty acid receptors in GLP-1 secretion remains to be clearly elucidated.

    The murine L-cell also expresses all three classes of PKCs (conventional, novel, and atypical) (R. Iakoubov, A. Izzo, A. Yeung, C.I. Whiteside, P.L.B., unpublished data). However, MUFA-induced GLP-1 secretion is mediated via activation of the atypical isoform PKC-ζ only (R. Iakoubov, A. Izzo, A. Yeung, C.I. Whiteside, P.L.B., unpublished data). The mechanism by which oleic acid activates PKC-ζ has yet to be determined, nor is it clear how oleic acid enters the cell. RT-PCR has confirmed the expression of fatty acid transport proteins, but their functional significance in MUFA-induced PKC-ζ activation and GLP-1 secretion is unclear. Further studies are clearly required to elucidate the exact mechanism of action of this important physiological regulator of GLP-1 release.


    As previously mentioned, it is not likely that glucose reaches the distal intestine to stimulate GLP-1 secretion, and in the physiological setting, GLP-1 secretion in response to ingested glucose is mediated indirectly by the vagus. Consistent with this hypothesis, GLP-1 release by isolated canine and rat L-cells is not directly affected by glucose (V.S.C. Wong, P.L.B., unpublished data; 62). Nonetheless, placement of glucose directly into the rat ileum has been found to stimulate the L-cell (27). Furthermore, murine L-cells do demonstrate dose-dependent GLP-1 secretion in response to glucose in vitro (40,41). Electrophysiological studies have shown that glucose effects a change in the membrane potential of the murine L-cell through closure of ATP-sensitive K+ (KATP) channels (40), as well as via depolarization due to co-transport of sodium and glucose through the sodium glucose transporter (41). Expression of the glucose sensor glucokinase has also been detected in the mouse intestinal L-cell in vivo (68). The opening of L-type Ca2+ channels is also associated with glucose treatment, which leads to GLP-1 secretion (40,55). Taken together, the electrogenic response of the murine L-cell to glucose is similar to the stimulus secretion coupling events that occur during glucose-stimulated insulin secretion, but unlike the β-cell, the absence of GLUT2 mRNA expression in the murine L-cell suggests that glucose transport is primarily facilitated by sodium glucose transporters. Nonetheless, a recent report has indicated that the KATP channel cannot be detected in the mouse L-cell in vivo (69) and KATP null mice do not show alterations in circulating GLP-1 levels (70). Thus, the physiological importance of the findings made with the murine L-cell in vitro remain to be confirmed.


    Leptin is a cytokine derived from adipocytes with potent effects on food intake (71). When administered to rats and mice in vivo, leptin demonstrates stimulatory effects on GLP-1 secretion, and these effects have also been observed in rodent and human L-cells in vitro (48). Furthermore, leptin resistance induced by a high-fat diet in mice is associated with reduced basal and nutrient-stimulated GLP-1 secretion. The leptin receptor is a product of the Ob-R gene, which is related to other class I cytokine receptors. Ob-R is spliced into five isoforms, but only the OB-Rb isoform, which is the long form of the receptor, has the necessary intracellular motifs for leptin signaling (71). After leptin binding to OB-Rb, janus kinase (JAK) phosphorylates residues on OB-Rb, which then serve as docking sites for signal transducer and activator of transcription (STAT) molecules (Fig. 5). Once phosphorylated, STAT molecules dimerize and enter the nucleus to mediate effects on gene transcription (71). Consistent with these findings, human and rodent L-cells express OB-Rb, and leptin treatment increases STAT-3 phosphorylation (48); however, it remains unknown as to how this results in enhanced GLP-1 release. Nonetheless, stimulatory effects of leptin have also been reported in STC-1 cells, such that cholecystokinin release occurs in response to leptin, and this appears to be dependent on p44/42 MAPK phosphorylation (72). Whether the effect of leptin on GLP-1 secretion from the L-cell is mediated by p44/42 MAPK remains to be determined. Nonetheless, these findings suggest a possible mechanism by which circulating GLP-1 levels are reduced in obese individuals (50).


    The insulin receptor is a tetrameric protein that consists of two extracellular α-subunits and two intracellular β-subunits linked together by disulfide bonds (71). Insulin binding to the α-subunits results in the phosphorylation of tyrosine residues in cytoplasmic domains of the β-subunits, thereby generating both tyrosine kinase activity and docking sites for a number of different intracellular proteins (Fig. 5). We have recently found that murine and human L-cells express the insulin receptor, and treatment of these cells with insulin results in insulin receptor, Akt, and p44/42 MAPK phosphorylation (49). Furthermore, insulin treatment stimulates GLP-1 secretion from the murine L-cell in a dose-dependent fashion. As Akt activation has recently been found to regulate catecholamine release from the chromaffin cell (73), these findings suggest one possible mechanism by which insulin stimulates GLP-1 release. Excitingly, both the murine and human L-cell were also found to exhibit insulin resistance after 24-h pretreatment with high concentrations of insulin. Furthermore, this insulin resistance was associated with altered basal and attenuated insulin-stimulated GLP-1 secretion, as well as with heterologous desensitization of the L-cell response to GIP (G.E.L., N. Flora, P.L.B., unpublished data; 49). When taken together, these findings suggest a possible mechanism by which GLP-1 levels are reduced in patients with type 2 diabetes (57), as well as in normal men in the lowest tertile of insulin sensitivity (25).

    Relevance to diabetes and insulin resistance.

    Numerous studies have now shown that GLP-1 levels after a meal are reduced in subjects with type 2 diabetes (57). These changes are not due to altered clearance of GLP-1 from the circulation (6) and appear to be independent of obesity (5). Although direct studies of the intestinal L-cell have implicated insulin resistance in this phenomenon (49), type 2 diabetes is associated with a broad range of metabolic disorders, and it therefore remains unclear as to the exact mechanism underlying the reductions in GLP-1 in this condition. Furthermore, because obesity is linked to leptin and insulin resistance, as well as to impaired secretion of GLP-1 (48), the coexistence of obesity and type 2 diabetes may result in further defects in the release of GLP-1. Although further studies are clearly required to elucidate the specific mechanisms leading to impaired GLP-1 secretion in both type 2 diabetes and obesity, several recent studies conducted in vivo have suggested that the insulin sensitizer metformin may increase circulating GLP-1 levels (74), possibly through effects on the L-cell. Detailed mechanistic studies of metformin action using in vitro models of the intestinal L-cell are clearly warranted.


    To date, only a limited number of studies have directly examined the signaling pathway used by the L-cell. Nonetheless, demonstration of the utility of GLP-1 as an antidiabetic agent has highlighted the importance of the intestinal L-cell as a potential target for pharmacological enhancement of GLP-1 levels. An improved understanding of the specific intracellular proteins that are crucial for GLP-1 secretion may lead to novel approaches to enhance GLP-1 levels in patients with type 2 diabetes.

    FIG. 1.
    FIG. 1.

    Tissue-specific posttranslational processing of proglucagon liberates different proglucagon-derived peptides. Prohormone convertase 2 in the α-cell releases glicentin-related pancreatic peptide (GRPP), glucagon, intervening peptide-1 (IP1), and the major proglucagon fragment. In the L-cell, cleavage of proglucagon by prohormone convertase 1/3 yields glicentin, oxyntomodulin, GLP-1, IP2, and GLP-2.

    FIG. 2.
    FIG. 2.

    Regulation of GLP-1 secretion by ingested nutrients. After a meal, nutrients in the duodenum activate a proximal-distal neuroendocrine loop, which stimulates GLP-1 secretion from L-cells in the ileum and colon. In rodents, GIP, released from K-cells, activates vagal afferents, which subsequently causes GLP-1 secretion through vagal efferents and enteric neurons that release acetylcholine (Ach) and GRP. Movement of nutrients toward more distal sections of the intestine leads to the direct interaction of nutrients with L-cells, which also stimulates GLP-1 secretion.

    FIG. 3.
    FIG. 3.

    Intracellular pathways of GLP-1 secretagogues activated by proximal nutrients. Binding of GRP or acetylcholine (Ach) to Gαq-linked GRP or M1 receptors, respectively, is associated with phospholipase C (PLC) activation and subsequent stimulation of conventional and novel isoforms of protein kinase C (c/n PKC). Activation of these receptors is also associated with increases in intracellular calcium and the phosphorylation of p44/42 mitogen-activated kinase. GIP receptor activation causes the stimulation of adenylyl cyclase (AC) via Gαs. This leads to an increase in cAMP and activation of PKA. The somatostatin receptor is coupled with Gαi, which inhibits AC, therefore inhibiting GLP-1 secretion. Binding of GABA to GABAA receptors depolarizes L-cells by channel opening and the efflux of chloride. Solid arrows = known pathways; dashed arrows = unknown pathways.

    FIG. 4.
    FIG. 4.

    Signaling components involved in direct nutrient-induced GLP-1 secretion. Glucose entry into L-cells via sodium glucose transporters (SGLT) causes an increase in ATP, which leads to KATP channel closure. Together with Na+ entry via sodium glucose transporters, this results in a change in membrane potential (ψ), which opens L-type voltage-gated calcium channels and leads to GLP-1 release. MUFA-induced GLP-1 secretion is mediated by GPR120 and is associated with increases in intracellular calcium and phosphorylation of Akt and p44/42 MAPK via Gαq. Alternatively, MUFAs may be transported via fatty acid transport proteins (FATP), leading to activation of PKC-ζ through an unknown mechanism to stimulate GLP-1 secretion. Solid arrows = known pathways; dashed arrows = unknown pathways.

    FIG. 5.
    FIG. 5.

    Signal transduction pathways of metabolic hormones that stimulate GLP-1 secretion. Insulin receptor activation results in phosphorylation of insulin receptor substrate (IRS) molecules and subsequent activation of Akt and p44/42 MAPK through phosphatidylinositol 3-kinase (PI3K) and mitogen-activated protein kinase kinase (MAPKK), respectively. Binding of leptin to its receptors in the L-cell results in STAT-3 phosphorylation, likely through janus kinase (JAK). Leptin receptor activation may also result in p44/42 MAPK phosphorylation. Solid arrows = known pathways; dashed arrows = unknown pathways.


    This work was supported by an operating grant from the Canadian Diabetes Association. G.E.L. was supported by graduate studentships from the Natural Sciences and Engineering Research Council of Canada and from the Banting and Best Diabetes Centre, University of Toronto. P.L.B. was supported by the Canada Research Chairs Program.


    • P.L.B. has received honoraria from and is a consultant for Amgen.

      This article is based on a presentation at a symposium. The symposium and the publication of this article were made possible by an unrestricted educational grant from Servier.

      The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

      • Accepted May 2, 2006.
      • Received March 23, 2006.


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    Sell Unused Diabetic Strips Today!

    Baked French Fries (Diabetes Friendly)

    By electricdiet / August 20, 2021

    Craving french fries? Try these easy baked french fries that are tossed in chili powder, cumin, and garlic, then roasted to crispy perfection!

    Bowl of baked French fries on table

    You may be wondering: can people with diabetes eat french fries?

    White potatoes aren’t the best choice from a glycemic index perspective, but for most of us, it doesn’t hurt to enjoy them every once in a while. Just be sure to monitor the total amount of carbs you’re eating.

    And these baked french fries are MUCH better for you than any french fries you’d get from a restaurant or fast-food chain! So when you need to squash a craving, this recipe is the way to go.

    These potato wedges are absolutely delicious. They’re coated in flavorful southwestern spices and roasted to crispy perfection — how great does that sound?

    But don’t just take my word for it. Whip up a batch and see for yourself!

    How to make baked french fries

    This recipe is easy to throw together for a tasty snack, appetizer, or side everyone will love!

    Potatoes, spices, and oil on a cutting board

    Step 1: Preheat the oven to 425°F.

    Step 2: Slice the potatoes into wedges, about 8 per potato, and put them in a gallon bag. Drizzle with olive oil.

    Step 3: Combine the chili powder, cumin, and garlic pepper in a small bowl, then add the mixture to the bag with the potatoes.

    Step 4: Seal the bag and squeeze it to move the wedges around until they are well-coated in the oil and spices.

    Zip-lock bag with potato wedges covered in oil and spices

    Step 5: Empty the bag of potatoes onto a baking sheet and spread them out so that none are touching.

    Step 6: Bake for 15 minutes, then flip the wedges and bake for another 10 minutes or until crispy.

    Baked French fries on a aluminum baking sheet

    Variations for this recipe

    There are a few ways you can get creative with these roasted potato wedges.

    I settled on a southwestern blend of spices because I was serving them alongside pulled pork tenderloin. But you’re welcome to use whatever spices you prefer.

    You can also cut the potato into different shapes than wedges. I’ve had success cutting them into rounds as well!


    Much like traditional french fries, roasted potato wedges are best served immediately. You want them while they’re hot and crispy!

    If you do have any leftovers, you can store them covered in the refrigerator for up to 5 days. I would recommend reheating them in the oven or a skillet.

    French fries on a cutting board

    Other tasty and healthy sides

    I love side dishes. Sometimes, they’re even better than the main course! If you’re looking for more tasty recipes to round out your meal, here are a few of my favorite side dishes:

    When you’ve tried this dish, please don’t forget to let me know how you liked it and rate the recipe in the comments below!

    Recipe Card

    Baked French Fries

    Baked French Fries

    Craving french fries? Try these easy baked french fries that are tossed in chili powder, cumin, and garlic, then roasted to crispy perfection!

    Prep Time:10 minutes

    Cook Time:25 minutes

    Total Time:35 minutes

    Author:Diabetic Foodie



    • Preheat the oven to 425°F.

    • Slice the potatoes into wedges, about 8 per potato, and put them in a zip-lock bag. Drizzle with olive oil.

    • Combine the chili powder, cumin, and garlic pepper in a small bowl, then add the mixture to the bag with the potatoes.

    • Seal the bag and squeeze it to move the wedges around until they are well-coated in the oil and spices.

    • Empty the bag of potatoes onto a baking sheet and spread them out so that none are touching.

    • Bake for 15 minutes, then flip the wedges and bake for another 10 minutes or until crispy.

    Recipe Notes

    This recipe is for 3 servings. Each serving is ¼ of a pound of potatoes (the number of potatoes in ¼ of a pound is going to depend on their size)
    Feel free to play around with the spice mixture or cut your potatoes into other shapes.
    Roasted potato wedges are best served immediately. Leftovers can be stored covered in the refrigerator for up to 5 days.

    Nutrition Info Per Serving

    Nutrition Facts

    Baked French Fries

    Amount Per Serving (0.25 lbs)

    Calories 117
    Calories from Fat 28

    % Daily Value*

    Fat 3.1g5%

    Saturated Fat 0.4g3%

    Trans Fat 0g

    Polyunsaturated Fat 0.4g

    Monounsaturated Fat 2.2g

    Cholesterol 0mg0%

    Sodium 92.2mg4%

    Potassium 496.7mg14%

    Carbohydrates 20.1g7%

    Fiber 0.2g1%

    Sugar 0g0%

    Protein 3.9g8%

    Vitamin A 200IU4%

    Vitamin C 0.8mg1%

    Calcium 60mg6%

    Iron 0mg0%

    Net carbs 19.9g

    * Percent Daily Values are based on a 2000 calorie diet.

    Course: Side Dish

    Cuisine: American

    Diet: Diabetic, Low Fat

    Keyword: french fries, potato wedges

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    Banana Oat Pancakes – My Bizzy Kitchen

    By electricdiet / August 18, 2021

    These banana oat pancakes could not be easier to make.  The blender does all the work!

    this is a photo of banana oat pancakes with blueberries

    How do you make fluffy oat pancakes?

    My friend Erica is the QUEEN of banana pancakes.  She uses one banana, one egg and a pinch of salt.  My first question to her was “where is the baking powder?”  That is the secret to fluffy pancakes with our without flour.  Simply let the batter sit overnight, or better yet, the night before for a quick breakfast.

    What do you need to make banana oat pancakes?

    • oats
    • banana
    • baking powder
    • salt
    • unsweetened almond milk (or any milk)
    • that’s it

    this is a photo of banana oat pancakes with blueberries

    What is the best sugar free pancake syrup?

    Being diabetic, I’ve literally tried every sugar free pancake syrup on the market.  While Smucker’s is up there, it is hit or miss if any store near me carries it.  Cary’s Sugar Free pancake syrup is my favorite, also the cheapest and only 5 calories and zero points for 1/4 cup.


    • 1/2 cup quick oats
    • 1 medium banana
    • 1 large egg
    • 2 teaspoons baking powder
    • 1 tablespoon almond milk
    • pinch of salt


    1. Blend ingredients in a blender. Let sit 30 minutes or overnight.
    2. Using 1/3 cup measuring for the batter, pan fry for 3-4 minutes a side over medium low heat.
    3. Optional: 1/2 cup frozen blueberries.


    On #wwpurple – this is zero points, on #wwblue it’s 2 points and on #wwgreen it’s 3 points.

    Nutrition Information:

    Yield: 2

    Serving Size: 1

    Amount Per Serving:

    Calories: 155Total Fat: 4gSaturated Fat: 1gTrans Fat: 0gUnsaturated Fat: 2gCholesterol: 93mgSodium: 635mgCarbohydrates: 26gFiber: 3gSugar: 8gProtein: 6g

    Did you make this recipe?

    Please leave a comment on the blog or share a photo on Instagram

    Don’t like bananas in your pancakes?  Try my cottage cheese blueberry pancakes – these are gluten free too!


    I am a widowed 52 year old trying to figure out my life after losing my husband after a long illness.

    Cooking and being in the kitchen feeds my soul!

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    Keto Chaffle Recipe | Diabetes Strong

    By electricdiet / August 16, 2021

    This keto chaffle recipe is ready in minutes and can be used in so many ways! Add sweet or savory toppings, use as low-carb bread for your favorite sandwich, and more.

    Finished chaffle from Keto Chaffle recipe, topped with bacon, fried egg, and scallions topped with bacon and a fried egg

    “Chaffles” have started a bit of a craze in the keto and low-carb community… and for a good reason! They’re only 2.7 net carbs per serving, and there are endless ways to put them to delicious use.

    Plus, you only need three ingredients and about 10 minutes for this keto chaffle recipe. You can enjoy them right away, or keep them in the freezer for super easy meal prep.

    How to make this keto chaffle recipe

    These tasty chaffles come together in just five simple steps. Let’s see how it’s done!

    Ingredients for recipe in separate bowls and ramekins, as seen from above

    Step 1: Preheat the waffle iron for 5 minutes. Spray with cooking oil once hot.

    Step 2: Mix together the eggs, cheese, and almond flour with a fork until well mixed.

    Eggs, cheese, and almond flour mixed together in a glass bowl

    Step 3: Pour half the mixture into the waffle maker until the surface is fully covered in a thin layer of batter.

    Step 4: Cook in the waffle machine for 4-5 minutes until crispy. When there is very little steam coming out, the waffle should be done.

    Finished chaffle in the waffle maker, as seen from above

    Step 5: Remove and set aside to cool. Repeat with the remaining batter.

    Two chaffles cooling on a wire rack, as seen from above

    How easy was that? This recipe will make two chaffles, but you can easily double, triple, or even quadruple the ingredients depending on how many you need!

    A finished chaffle on a white plate, ready for toppings

    What toppings to serve with your chaffle

    When it comes to toppings or ways to enjoy your chaffles, the possibilities are truly endless. Even though we use cheese in this recipe, the final flavor turns out pretty neutral. That means you can enjoy them sweet, savory, or however you like!

    One of the most obvious options is to use them exactly like regular waffles. Add your favorite low-carb toppings, like sugar-free maple syrup or fresh fruit, and dig in. I personally love adding some shredded coconut and sugar-free chocolate chips.

    You can also use chaffles as a low-carb substitute for toast. Top with bacon and a fried egg for a delicious open-faced breakfast sandwich.

    What about a quick lunch? You can layer up with your favorite deli meats, cheese, and condiments, or make yourself a keto chicken salad sandwich!

    From breakfast to dessert, chaffles are so versatile. You’ll never stop finding yummy ways to use them.

    Chaffled topped with bacon, a fried egg, and scallions


    Chaffles are a great option for simple meal prepping. I love having them in my freezer and ready to use at a moment’s notice!

    Once the chaffles have finished cooking and fully cooled, you can freeze them in an airtight container for up to 2 weeks. When you’re ready to enjoy, reheat your chaffles in the toaster or a preheated waffle iron to quickly crisp them up before serving.

    A fork holding a bite of chaffle and toppings in front of white space

    Other delicious keto-friendly recipes

    So many comfort foods can be adapted for a low-carb diet. You just have to know how to make a few ingredient swaps! Here are a few of my favorite foods with keto-friendly updates that I know you’ll love:

    Looking for more low-carb ways to satisfy your sweet tooth? Make sure to check out my roundup of the best keto-friendly dessert recipes!

    When you’ve tried this dish, please don’t forget to let me know how you liked it and rate the recipe in the comments below!

    Recipe Card

    Keto Chaffle Recipe

    This keto chaffle recipe is ready in minutes and can be used in so many ways! Add sweet or savory toppings, use as low-carb bread for your favorite sandwich, and more.

    Prep Time:5 minutes

    Cook Time:5 minutes

    Total Time:10 minutes


    Finished chaffle from Keto Chaffle recipe, topped with bacon, fried egg, and scallions


    • Preheat the waffle iron for 5 minutes. Spray with cooking oil once hot.

    • Mix together the eggs, cheese, and almond flour with a fork until well mixed.

    • Pour half the mixture into the waffle maker until the surface is fully covered in a thin layer of batter.

    • Cook in the waffle machine for 4-5 minutes until crispy. When there is very little steam coming out, the waffle should be done.

    • Remove and set aside to cool. Repeat with the remaining batter.

    Recipe Notes

    This recipe is for 2 chaffles. Each chaffle is 1 serving.
    To store, allow the chaffles to cool fully after cooking, then freeze in an airtight container for up to 2 weeks.
    To reheat, place in the toaster or a preheated waffle iron to quickly crisp them up before serving.

    Nutrition Info Per Serving

    Nutrition Facts

    Keto Chaffle Recipe

    Amount Per Serving (1 chaffle)

    Calories 270
    Calories from Fat 185

    % Daily Value*

    Fat 20.5g32%

    Saturated Fat 8.8g44%

    Trans Fat 0g

    Polyunsaturated Fat 0g

    Monounsaturated Fat 0g

    Cholesterol 225mg75%

    Sodium 410mg17%

    Potassium 70mg2%

    Carbohydrates 3.5g1%

    Fiber 0.8g3%

    Sugar 0.3g0%

    Protein 21.5g43%

    Net carbs 2.7g

    * Percent Daily Values are based on a 2000 calorie diet.

    Course: Breakfast, Dessert, Side Dish, Snack

    Cuisine: American

    Diet: Diabetic, Gluten Free

    Keyword: Chaffle, chaffle recipe, gluten-free, Keto chaffle

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    Delicious Dinner Ideas for Diabetics with Quick Diabetic Dinner Recipes

    By electricdiet / August 14, 2021

    Easy Dinner Ideas for Diabetics

    Does it feel like you are always looking for dinner ideas for diabetics? Our goal is to make eating healthy easy and mainstream! Of course, Team Holly always have quick diabetic recipes because who has time to cook these days? Even cooking food approved by the American Diabetes Association guidelines can be delicious with Holly Clegg’s trim and terrific recipes like this Smothered Chicken.  That’s why Holly decided to highlight easy diabetic recipes in all her cookbooks.  So, look for a “D” in cookbooks: Kitchen 101, Eating Well To Fight Arthritis and Eating Well Through Cancer to quickly find diabetic recipes.

    Why Protein for Type 2 Diabetes

    The most common form of Diabetes Mellitus is Type 2 Diabetes, which occurs when the body does not appropriately respond to and produce insulin; which results in high blood sugar. Protein from meats such as poultry does not contain carbohydrate (which turns into glucose, aka sugar in the blood) so they do not raise blood sugar levels.

    Low Fat Protein for Heart Health

    Choose protein low in saturated fat is important for heart health and especially for those with diabetes. Choose poultry without the skin for less saturated fat and cholesterol. Trim and terrific healthy easy recipes make excellent dinner ideas for diabetics such as this Smothered Chicken recipe from KITCHEN 101: Secrets to Cooking Confidence using boneless skinless chicken breasts. Who says you can’t have your favorite foods while still eating healthy and keeping to your diabetic meal plan?

    Fiber for Blood Sugar Control

    Be sure to choose brown rice in your dinner ideas for diabetics as 1 cup of brown rice as 3 times the fiber as white rice. Fiber helps control blood sugar levels from spiking in those with Type 2 diabetes. Fiber also helps in weight management because it aides in digestion and helps keep you full and satisfies hunger.

    Why I Use Kitchen Scissors

    Do you have kitchen scissors? If not, kitchen scissors are truly one of our favorite gadgets. We always use them to cut chicken into tenders or to trim chicken. When you are making any diabetic chicken recipes, you can buy chicken breasts and cut into tenders yourself.

    These kitchen scissors are dishwasher safe and any gadget that helps cut kitchen time is high on our list. You’ll find so many uses for kitchen scissors from trimming chicken to cutting pizza.

    Most Popular Glazed Salmon Recipe Tops Dinner Ideas for Diabetics!

    Salmon is a great choice for a lean protein and healthy fish.   Glazed Salmon (another KITCHEN 101 recipe) is one of most popular recipes and it even went viral on YouTube!! Wait until you try this simple salmon recipe!  Remember, when you use trim and terrific recipes you will be eating healthy and a diabetic diet is really the healthiest way to eat.  You also can reduce your risk of diabetes by cooking at home–so why not?

    Cookbooks Highlight Quick Diabetic Recipes!

    Best of all, you probably have all the ingredients in your pantry and these books include all your favorite recipes–yes, and they are great diabetic dinner ideas! Our go-to amazing Peanut Butter Cookies recipe in KITCHEN 101 with 5 ingredients is so good!

    Get All of Holly’s Healthy Easy Cookbooks

    The post Delicious Dinner Ideas for Diabetics with Quick Diabetic Dinner Recipes appeared first on The Healthy Cooking Blog.

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    Metabolic Endotoxemia Initiates Obesity and Insulin Resistance

    By electricdiet / August 12, 2021


    Diabetes and obesity are two metabolic diseases characterized by insulin resistance and a low-grade inflammation. Seeking an inflammatory factor causative of the onset of insulin resistance, obesity, and diabetes, we have identified bacterial lipopolysaccharide (LPS) as a triggering factor. We found that normal endotoxemia increased or decreased during the fed or fasted state, respectively, on a nutritional basis and that a 4-week high-fat diet chronically increased plasma LPS concentration two to three times, a threshold that we have defined as metabolic endotoxemia. Importantly, a high-fat diet increased the proportion of an LPS-containing microbiota in the gut. When metabolic endotoxemia was induced for 4 weeks in mice through continuous subcutaneous infusion of LPS, fasted glycemia and insulinemia and whole-body, liver, and adipose tissue weight gain were increased to a similar extent as in high-fat–fed mice. In addition, adipose tissue F4/80-positive cells and markers of inflammation, and liver triglyceride content, were increased. Furthermore, liver, but not whole-body, insulin resistance was detected in LPS-infused mice. CD14 mutant mice resisted most of the LPS and high-fat diet–induced features of metabolic diseases. This new finding demonstrates that metabolic endotoxemia dysregulates the inflammatory tone and triggers body weight gain and diabetes. We conclude that the LPS/CD14 system sets the tone of insulin sensitivity and the onset of diabetes and obesity. Lowering plasma LPS concentration could be a potent strategy for the control of metabolic diseases.

    The outbreak of a fat-enriched diet in Western countries is becoming a problem of the utmost importance. Obesity is the result of a complex interaction between genetic and environmental factors. Among the latter, changes in eating habits to increase fat intake are involved in the increased occurrence of metabolic diseases, such as obesity and diabetes, which are bearing features of the metabolic syndrome. The major metabolic consequence of a high-fat diet is that insulin action and the regulatory mechanisms of body weight are impaired through a well-described lipotoxic effect (1). In addition, it has been recently determined that obesity and insulin resistance are associated with low-grade chronic systemic inflammation (2). In models of diet-induced and genetic obesity, the adipose tissue presents increased expression and content of proinflammatory cytokines such as tumor necrosis factor (TNF)-α (3,4), interleukin (IL)-1 (3,4), and IL-6 (4). This cytokine production is then deleterious for muscle insulin action; for example, TNF-α has been shown to cause insulin resistance by increasing serine phosphorylation on insulin receptor substrate-1 (5), leading to its inactivation. The consequent insulin resistance will favor hyperinsulinemia and excessive hepatic and adipose tissue lipid storage. However, while extensive research is dedicated to the effects of an inflammatory reaction on energy metabolism, the triggering factor linking inflammation to high-fat diet–induced metabolic syndrome remains to be determined. Recently, it has been shown that nutritional fatty acids activate toll-like receptor-4 (TLR4) signaling in adipocytes and macrophages and that the capacity of fatty acids to induce inflammatory signaling in adipose cells or tissue and macrophages is blunted in the absence of TLR4 (6). Furthermore, adipose tissue lipolysis, from hypertrophied adipocytes, could serve as a naturally occurring ligand for TLR4 to induce inflammation (7). In addition, TLR4 mRNA concentration was induced during adipocyte differentiation, further enhancing free fatty acid–induced inflammation (8).

    A very encouraging and innovative hypothesis has recently been proposed whereby the microbial ecology in humans could be an important factor affecting energy homeostasis (i.e., individuals predisposed to obesity may have gut microbial communities that would favor the occurrence of the metabolic diseases) (911). Although the proposed mechanism was to promote a more efficient extraction and/or storage of energy from a given diet, the impact of microbiota on the occurrence of a low-tone inflammatory status was not determined. However, another study recently showed that antibiotic treatment partially protects against type 1 diabetes in a diabetes-prone rat developing insulitis (12). The authors proposed that altering the gut microbiota composition by antibiotic treatment had reduced the antigenic load and hence the inflammatory reaction that had potently led to pancreatic β-cell destruction. In that respect, we have been looking for a factor of microbial origin that would trigger and maintain a low-tone continuous inflammatory state when feeding on a high-fat diet. We hypothesized that the bacterial lipopolysaccharide (LPS) from the Gram-negative intestinal microbiota would fulfill all the prerequisites to be eligible. Thus, endogenous LPS is 1) continuously produced in the gut by the death of Gram-negative bacteria and physiologically translocated into intestinal capillaries through a TLR4-dependent mechanism (13); 2) transported from the intestine toward target tissues by a mechanism facilitated by lipoproteins, notably chylomicrons freshly synthesized from epithelial intestinal cells in response to a high-fat diet (14); and 3) triggers the secretion of proinflammatory cytokines when it binds to the complex of mCD14 and the TLR4 at the surface of innate immune cells (15,16). Therefore, we aimed to demonstrate that LPS would be an early factor in the triggering of high-fat diet–induced metabolic diseases.


    Twelve-week-old male C57bl6/J mice (Charles River, Brussels, Belgium) and CD14 mutant male mice bred in a C57bl6 background (Jackson Laboratory, Bar Harbor, ME) were housed in a controlled environment (inverted 12-h daylight cycle, lights off at 10:00 a.m.) with free access to food and water. All of the following animal experimental procedures were validated by the local ethical committee of the Rangueil Hospital and by the Université catholique de Louvain Animal Ethical Committee.

    Mice were fed a control (A04, Villemoisson sur Orge, France) or a high-fat, carbohydrate-free diet for 2 or 4 weeks following protocols. The diet contained 72% fat (corn oil and lard), 28% protein, and <1% carbohydrate as energy content (17). In a subset of mice, obesity and diabetes were induced by a high-fat diet and mice killed after 4 or 24 weeks.

    Energy intake measurements.

    Energy intake was recorded twice weekly for 4 weeks, two mice per cage. Pellets and spillage were weighed separately. The mean value for the weekly assessment was calculated. This method was performed similarly for all assessments.

    Surgical procedures, infusions, and isotope measurements.

    Mice were implanted subcutaneously with an osmotic minipump (Alzet Model 2004; Alza, Palo Alto, CA) as previously described (17). The pumps were filled either with NaCl (0.9%) or LPS (from Escherichia coli 055:B5; Sigma, St. Louis, MO) to infuse 300 μg · kg−1 · day−1 for 4 weeks. For the clamp studies, an intrafemoral catheter was indwelled as previously described (18). Insulin sensitivity was assessed by the euglycemic-hyperinsulinemic clamp as described (19). Briefly, 6-h–fasted mice were infused with insulin at a rate of 18 (pharmacological, to assess whole-body insulin sensitivity) or 1 (physiological, to assess the effect of insulin on the inhibition of endogenous glucose production) mU · kg−1 · min−1 for 3 h. For glucose turnover measurements, d-(3H)3-glucose (Perkin Elmer, Boston, MA) was simultaneously infused at rate of 30 μCi/kg, as described (17). d-(3H-3)-glucose enrichments were determined from total blood after deproteinization by a Zn(OH)2 precipitate, as described (19). To assess whether CD14 mice were sensitive to LPS-induced inflammation, fasted mice were continuously infused with LPS (0.5 mg · kg−1 · h−1) for 3 h. Liver and white adipose tissue were collected and frozen at −80°C.

    Glucose and LPS tolerance tests.

    Intraperitoneal or oral glucose tolerance tests were performed as follows: 6-h–fasted mice were injected with glucose into the peritoneal cavity (1 g/kg glucose, 20% glucose solution) or by gavage (3 g/kg glucose, 66% glucose solution). Blood glucose was determined with a glucose meter (Roche Diagnostics) on 3.5 μl blood collected from the tip of the tail vein. A total of 20 μl blood was sampled 30 min before and 15 or 30 min after the glucose load to assess plasma insulin concentration. An LPS tolerance test was performed as follows. Fasted mice were gavaged by LPS (300 μg/kg) diluted in water or corn oil (100 μl) or without LPS. Blood was collected from the tip of the tail vein before and 30 min after gavage. Plasma was separated and frozen.

    Microbial enumeration in intestinal content by fluorescent in situ hybridization.

    The caecal contents collected post mortem from mice were stored at −80°C. Samples were thawed on ice and diluted 1:10 in sterile ice-cold 0.1 mol/l PBS, pH 7.0. The suspension was homogenized by pipetting and centrifuged at 3,500g for 15 min to remove particulate matter. The supernatant containing the bacterial cells was fixed overnight in 4% (wt/vol) paraformaldehyde. The bacterial cells were then washed and resuspended twice in sterile PBS and finally stored in 50% ethanol in PBS at −20°C until hybridization with appropriate molecular probes targeting specific regions of 16S rRNA. The probes used were EREC 482, Bac303, MIB 661, Lab158, Bif164, SRB 687, and probe D specific for the Eubacterium rectale/Clostridium coccoides group, Bacteroides species, mouse intestinal bacteria, Lactobacilli/Enterococci, Bifidobacterium species, sulfate-reducing bacteria, and the Enterobacteriaceae, respectively. The nucleic acid stain DAPI (4″6-diamidino-2-phenylindole) was used for total bacterial counts. The DNA probes were tagged with the Cy3 fluorescence dye so that the hybridized samples could be examined using fluorescence microscopy, as described previously (20). Results were expressed as log10 (bacterial cells per gram caecal content wet weight).

    Real-time quantitative PCR.

    Total RNAs from liver, muscle, and white adipose tissue were prepared using TriPure reagent (Roche, Basel, Switzerland) as described (21). PCRs were performed using an ABI PRISM 5700 Sequence Detection System instrument and software (Applied Biosystems, Foster City, CA) as described (21). Primer sequences for the targeted mouse genes are available upon request (E-mail: patrice.cani{at}

    Western blots.

    One hundred milligrams of liver were homogenized in lysis buffer, and the proteins were separated onto a polyacrylamide gel and transferred to polyvinylidine fluoride membrane, as described (17). The membranes were incubated overnight at 4°C with the indicated antibody of nuclear factor κB-p65 (Cell signaling, Saint Quentin Yvelines, France) and inhibitor of κB kinase β-P (Santa Cruz, Le Perray en Yvelines, France). The binding of the specific primary antibody was quantified as described (17).

    Adipose tissue morphometry and F4/80 staining.

    The mean relative proportion of adipocytes was estimated by a point-counting technique, on paraffin-embedded hematoxilin eosine staining counterstained sections of subcutaneous tissue. The number of adipocytes per microscopical field (density) was determined at a magnification of ×200. Total count ranged from 2,300 to 6,600 cells per condition. The mean surface area of the adipocytes (μm2) was calculated using image analyzer software (Visilog 6, Courtaboeuf, France). Each adipocyte was manually delineated, and 700–1,000 adipocytes per condition were assessed. F4/80 staining was performed as follows. Ethanol-fixed, paraffin-embedded adipose tissue sections were deparaffinized and rehydrated. Sections were blocked in normal serum and incubated overnight with primary rat anti-mouse F4/80 monoclonal antibody (1/1,000; Serotec, Oxford, U.K.). Endogenous horseradish peroxidase activity was quenched with incubation with 3% hydrogen peroxide for 20 min. Secondary antibody staining was performed using goat anti-rat biotinylated Ig Ab (1/500, 30 min, room temperature) and streptavidin horseradish peroxidase conjugated (1/500, 30 min, room temperature) and detected with 3,3′-diaminobenzidine. Sections were counterstained with hematoxylin before dehydration and coverslip placement. The number of F4/80-positive cells per microscopical field was calculated and divided by the total number of adipocytes in sections. Twelve to 17 fields were counted per condition.

    Liver histology.

    A fraction of the main liver lobe was fixed-frozen in Tissue-tek in liquid nitrogen–cold isopentane. For the detection of neutral lipids, frozen sections were sliced and stained with the oil red O, using 0.5% oil red O dissolved in propylene glycol for 10 min at 60°C. The sliced sections were then counterstained.

    Biochemical analyses.

    Plasma LPS determinations in the mouse were performed using a kit based upon a Limulus amaebocyte extract (LAL kit; Cambrex BioScience, Walkersville, MD). Samples were diluted 1/40 to 1/80 and heated during 10 min at 70°C. Internal control of recovery calculation was included in the assessment. Plasma insulin concentrations were determined in 5 μl of plasma using an enzyme-linked immunosorbent assay kit (Mercodia, Upssala, Sweden), following manufacturer’s instructions. Liver triglyceride was determined as follows: total neutral lipids were extracted from 30 to 50 mg liver in CHCl3-MeOH (2:1). The organic extracts were air dried, reconstituted in isopropanol, and assayed for triglycerides by measuring the glycerol produced after enzymatic hydrolysis of triglycerides using a commercial kit (Triglycerides GPO Trinder; Sigma, Lyon, France).

    Statistical analysis.

    Results are presented as means ± SE. Statistical significance of differences was analyzed by one-way ANOVA followed by post hoc (Bonferroni’s multiple comparison test) using GraphPad Prism version 4.00 for Windows (GraphPad Software, San Diego, CA). Data with different superscript letters and * or § are significantly different at P < 0.05, according to the post hoc ANOVA statistical analysis.


    High-fat diet increased endotoxemia.

    Because LPS is an important component of lipoprotein (14), and dyslipidemia an important feature of metabolic disease, we studied the high-fat diet–fed mouse to determine the effect of a high-fat diet on the occurrence of plasma LPS concentration. We determined whether plasma LPS could be a physiological regulating factor dependent on the feeding status and measured the endotoxemia throughout the day. Our data show the existence of diurnal variations of plasma LPS concentration, which reaches a zenith at the end of the dark, feeding period (Fig. 1A). Furthermore, a short time (4 weeks) of high-fat feeding caused a disruption of this cycle, with endotoxemia remaining high throughout the whole day. This increase was still 10–50 times lower than values that could be reached during septicemia or other infections (22). Hence, we defined it as a metabolic endotoxemia. To determine whether a high-fat diet could bring about a modification within the intestinal microbiota, which might account for the observed increased levels of plasma LPS and, thus, metabolic endotoxemia, we characterized and quantified the main bacterial families of cecal microbiota in high-fat–fed and control mice. Maintenance of the mice on a high-fat diet for 4 weeks resulted in a significant modulation of dominant bacterial populations within the intestinal microbiota. Numbers of the newly recognized Gram-negative operating taxonomic unit, Bacteroides-like mouse intestinal bacteria (covered by the MIB661 probe), which reside within the Cytophaga-Flavobacter-Bacteroides phylum, were significantly reduced in animals fed the high-fat diet. These bacteria are closely related to the Bacteroides-Prevotella group but are not enumerated by the Bacteroides probe Bac303. Bacteria enumerated by the MIB661 probe are the most numerous Gram-negative bacteria within the mouse intestinal tract and, together with the Eubacterium rectale-Clostridium coccoides group, constitute the dominant members of the mouse intestinal microbiota (23). Numbers of the Eubacterium rectale-Clostridium coccoides group and bifidobacteria were also significantly lower in animals fed the high-fat diet compared with controls (Fig. 1B). Interestingly, these bacterial families are predominantly Gram positive. Second, we assessed whether exogenous LPS could be absorbed by a high-fat diet. We orally administered LPS diluted in oil or water to mice and found that endotoxemia rose after the LPS-oil mixture administration only (Fig. 1C). In addition, administration of oil alone also tended to increase endotoxemia. Therefore, both sets of experiments suggested that exogenous and endogenous LPS origin participate toward increased endotoxemia in the presence of a lipid-rich diet.

    Chronic experimental metabolic endotoxemia induces obesity, diabetes, and liver insulin resistance.

    Therefore, to causally link high-fat diet–increased plasma LPS concentration to metabolic disease, we mimicked the metabolic endotoxemia obtained during a 4-week high-fat feeding by implanting a subcutaneous osmotic minipump and performing a chronic continuous subcutaneous infusion of LPS for 1 month (Fig. 2A). A 72% high-fat–enriched diet increased endotoxemia 2.7-fold when compared with control fed mice. Interestingly, this increase was modest 1.4-fold when the mice were fed with a 40% high-fat–enriched diet. Therefore, we performed all studies with the 72% high-fat–enriched diet. Fasted glycemia was higher in chronic LPS–infused compared with saline-infused mice (Fig. 2B). Similarly, 15 min following oral glucose challenge, blood glucose concentration was higher in LPS- than in saline-treated mice (Fig. 2B). Similar data were obtained in 4-week high-fat diet–fed mice for the fasted and 15-min time points (Fig. 2B). However, the blood glucose concentration remained higher in the high-fat diet–fed mice compared with control and LPS-infused mice. The area under curves were higher in the LPS and high-fat diet–fed mice when compared with control mice (Fig. 2B inset). Furthermore, fasted insulinemia was higher in LPS- and high-fat–treated mice than in control mice. However, glucose-induced insulin secretion was normal in the chronic LPS-infused animals and was lower than control mice in high-fat diet–fed mice (Fig. 2C). Increased fasted plasma glucose and insulin levels were associated with liver but not with whole-body insulin resistance in chronic LPS–infused, whereas high-fat diet–fed mice were characterized with whole-body, but not liver, insulin resistance (Fig. 2D).

    In addition, the 4-week LPS infusion increased body weight to the same extent as a 4-week high-fat diet regimen (Fig. 2E). The visceral and subcutaneous adipose depots were similarly increased (Fig. 2F). This increase was more pronounced in the high-fat diet–fed mice. Liver weight was higher in the LPS-infused than in control and high-fat diet–fed mice (Fig. 2G). Liver triglyceride content was increased in both LPS-infused and high-fat diet–fed mice when compared with control mice, whereas significance was reached in the high-fat diet–fed group only (Fig. 2H). Energy intakes were quantified and found to be increased in the high-fat diet–fed group only when compared with control mice (Fig. 2I).

    Metabolic endotoxemia triggers the expression of inflammatory factors similarly to high-fat diet: a CD14-dependent mechanism.

    We determined the expression pattern of the main inflammatory factors involved in metabolic diseases, after 2 and 4 weeks of a high-fat diet or subcutaneous chronic LPS infusion. The mRNA concentrations of TNF-α, IL-1, IL-6, and plasminogen activator inhibitor (PAI)-1 were generally increased in various combinations in the liver, as well as in the visceral and subcutaneous depots, and in muscle of high-fat diet–fed and LPS-infused mice (Fig. 3). Interestingly, liver cytokines were increased as early as 2 weeks after commencement of the diet (Fig. 3A), which could also be related to the early augmentation of plasma LPS concentration (3.4 ± 0.3 EU/ml, compared with Fig. 2A). From these data, we can conclude that LPS mimics, to some extent, high-fat diet–induced inflammation and could be responsible for impaired metabolism.

    To demonstrate the causal link between LPS and obesity/diabetes, we studied CD14 mutant mice. In wild-type (WT) mice, the acute 3-h intravenous LPS infusion markedly increased IL-6, PAI-1, and IL-1 mRNA concentrations in the subcutaneous adipose depot, whereas this increase was dramatically blunted in CD14 mutant mice (Figs. 4A and B). Furthermore, the concentration of phosphorylated nuclear factor-κB and IKK forms were increased in the liver of WT mice and unchanged in CD14 mutant mice (Fig. 4C). We then quantified the effect of the chronic LPS infusion in WT and CD14 mutant mice. The data showed that body weight gain and visceral and subcutaneous adipose depot weights were increased in WT but unchanged in CD14 mutant mice (Figs. 5A and B). Furthermore, fasted and glucose-stimulated glycemias were augmented in WT-LPS–infused mice when compared with WT-CT mice (Fig. 5C). This observation was not apparent when CD14 mutant mice were infused with LPS. The area under the curve was increased in response to LPS infusion in the WT mice only (Fig. 5C, inset). Plasma insulin concentrations were similar in the basal and glucose-stimulated conditions for all groups (Fig. 5D). Chronic LPS infusion increased liver weight in the WT mice only (Fig. 5E). Triglycerides were increased by 30% in the liver of LPS-infused WT mice but did not reach statistical significance when compared with all other conditions (Fig. 5F). All together, these data confirm that LPS is directly implicated in the alterations of body weight and glucose metabolism observed in Figs. 1 and 2 in the previous experiments.

    CD14: the main LPS receptor sets the tone of insulin sensitivity and the occurrence of obesity and diabetes.

    We found that CD14 mutant mice are hypersensitive to insulin, as determined by the euglycemic-hyperinsulinemic clamp (Fig. 6A). These data show that CD14 itself can affect the basal physiological state of insulin sensitivity. Therefore, we challenged the mutant mice with a high-fat diet to delineate the relevance of hypersensitivity to insulin. We found that hypersensitivity to insulin allows the mutant mice to delay, without totally preventing, the occurrence of pathological insulin resistance and increased body weight (Fig. 6B) when fed a high-fat diet. Liver weight progressively increased and reached statistical significance after 24 weeks of a high-fat diet in WT mice (Fig. 6C). This increase was prevented in CD14 mutant mice. Similarly, liver triglyceride concentration was increased after only 4 weeks of a high-fat regimen (Fig. 6D). This was totally blunted in the CD14 mutant mice. Similarly, oil red O staining showed that lipid accumulation was blunted in the CD14 mutant fed a high-fat diet or infused with LPS (Fig. 6E). WT and CD14 mutant mice both became glucose intolerant after 24 weeks of high-fat diet (Fig. 6F). In addition, WT high-fat diet–fed (WT-HF) but not CD14 mice became hyperinsulinemic after 24 weeks of a high-fat diet (Fig. 6G).

    We performed a closer analysis of these mice after 4 weeks of a high-fat diet, when insulin resistance is already present in WT but not in CD14 mutant mice. After only 4 weeks of high-fat diet, the CD14 mutant mice had normal fasted glycemia, remained normally tolerant to glucose (Fig. 7A), and did not become hyperinsulinemic (Fig. 7B) in the fasting state when compared with WT mice. Visceral adipose depot weight was increased in WT but not in CD14 mutant mice fed a high-fat diet (Fig. 7C), whereas subcutaneous adipose depot was increased in both genotypes during high-fat diet (Fig. 7C). Furthermore, TNF-α and PAI-1 mRNA concentrations were both increased by the high-fat diet in WT mice, whereas this increase was totally blunted in the CD14 mutant mice (Fig. 7D). In the liver, a similar tendency was observed as in the adipose depot, showing that in absence of CD14 a high-fat diet could not induce inflammation (Fig. 7E). Although, PAI-1 mRNA levels are significantly elevated in CD14 high-fat–fed mice relative to controls.

    CD14 mutant mice resist high-fat diet–induced adipose depot macrophages infiltration and partly reduce adipocyte hypertrophy.

    Adipose cell size was increased in WT-HF mice when compared with WT-CT mice and to a lower extent in CD14 mutant mice (Figs. 8A, C, and D). Conversely, the mean adipocyte size was reduced in WT-LPS infused but not in CD14 mutant mice (Figs. 8BD). These changes in adipocyte size were accompanied with variations in F4/80-positive cells present in the subcutaneous adipose depots (Fig. 8E). High-fat feeding and LPS increased the number of F4/80-positive cells in WT mice. This was totally abolished in CD14 mutant mice (Fig. 8E).


    We report here for the first time that high-fat feeding augments plasma LPS at a concentration sufficient to increase body weight, fasted glycemia, and inflammation. LPS infusion in normal diet–fed mice causes a metabolic response similar, to some extent, to high-fat feeding. LPS receptor–deleted mice (i.e., CD14 mutants) are hypersensitive to insulin, and the occurrence of insulin resistance, obesity, and diabetes is delayed in response to high-fat feeding. Hence, we conclude that the LPS/CD14 system sets the threshold at which high-fat diet–induced metabolic disease occurs.

    Central to metabolic diseases is insulin resistance associated with a low-grade inflammatory status. In our quest to determine a triggering factor of the early development of metabolic disease, we looked for a molecule involved early in the cascade of inflammation and identified LPS as a good candidate. Furthermore, LPS is a strong stimulatory of the release of several cytokines that are key inducers of insulin resistance. The concept of dietary excess is essentially linked to high-fat feeding–induced inflammation (24). As we identified here, LPS as a putative factor for the triggering of metabolic diseases. We needed to challenge our hypothesis within a pathological context. We fed mice a high-fat diet for a short period of 2–4 weeks and showed that high-fat diet increases the circulating concentration of plasma LPS. Hence, we defined this increase in plasma LPS concentration induced by high-fat feeding as metabolic endotoxemia. It is noteworthy that the endotoxemia reached was 10–50 times lower than that obtained during septic shock (22,25). The mechanisms allowing enteric LPS absorption are unclear but could be related to an increased filtration of plasma LPS into lymph with fat absorption (26). Endogenous LPS is continually produced within the gut by the death of Gram-negative bacteria (27) and is absorbed into intestinal capillaries (28,29) to be transported by lipoproteins (14,15). Therefore, we assessed whether a high-fat diet changed intestinal microbiota and showed that among the dominant members of the intestine the Gram-negative Bacteroides-related bacteria MIB were significantly reduced in the high-fat diet–fed animals compared with control animals. However, numbers of the dominant Gram-positive group, the Eu. rectale-Cl. coccoides group, were also reduced, as were numbers of bifidobacteria, a group of bacteria that have been shown to reduce intestinal endotoxin (LPS) levels in mice and improve mucosal barrier function (30,31). Supporting our conclusion, apart from further evidence from the literature, was the result that an acute ingestion of LPS, diluted in oil, reproduced high-fat diet–increased plasma LPS concentration. Furthermore, oil by itself can acutely increase endotoxemia. We could therefore suggest that plasma LPS levels depends on feeding status and are physiologically regulated nutrients. We challenged this hypothesis in mice assessing endotoxemia throughout the day and showed that plasma LPS concentration increases progressively over the feeding period (night in the mouse). As the daily endotoxemia cycle was totally disrupted during high-fat feeding, our data showed that the fat content in food is an important regulator of plasma LPS concentration. In light of the changes observed in the microbiota of high-fat diet–fed mice, one could suggest that an increased continuous intestinal absorption of LPS could maintain steady and elevated the endotoxemia. Such a conclusion is also supported by our epidemiological study in humans in which healthy patients feeding on a fat-enriched diet were characterized by a higher fasting endotoxemia (data not shown).

    To assess whether a two- to three-times–increased daily endotoxemia could be considered as a physiological regulator, glucose metabolism was assessed in mice in which we continuously infused a very low rate of LPS. We found that the body and adipose depot weights and fasted glycemia were increased to the same extent as during high-fat feeding. Furthermore, the chronic LPS infusion induced liver insulin resistance and was associated with fasted hyperinsulinemia. As whole-body insulin resistance was not induced by the LPS infusion, our data showed the liver as a first target of LPS-induced insulin resistance. The role of LPS as a regulator of energy metabolism has been proposed previously (32,33). In most of the studies that described the anorectic and metabolic effects of LPS, endotoxemia was very elevated or given as single shot and witnessed a state of acute phase infection. Here, we showed that the chronic infusion of a very low rate of LPS increased body weight. However, this increase was not due to an excessive energy intake.

    We then studied the CD14 mutant mice, in which the expression of the protein is suppressed by a point mutation (34). CD14 is a key molecule in innate immunity (35). It is a multifunctional receptor phosphatidylinositol phosphate–anchored glycocoprotein of 55 kDa constitutively expressed in considerable amounts on the surface (mCD14) of mature monocytes, macrophages, and neutrophils (36,37). The binding of LPS to the complex of mCD14 and TLR4 at the surface of the innate immune cells (15) triggers the secretion of proinflammatory cytokines (16), consequently affecting insulin action. Mice depleted of the CD14 gene lack the innate immune response to bacterial LPS, and CD14-deficient macrophages do not secrete proinflammatory cytokines when stimulated with LPS (38). We have shown here that CD14 mutant mice fed a normal diet were hypersensitive to insulin. Therefore, CD14 could clearly be an early modulator of insulin sensitivity and, indeed, when challenged by a high-fat diet for 1 month the insulin-hypersensitive CD14 mutant mice did not become insulin resistant, glucose intolerant, or diabetic. However, after long-term feeding (24 weeks), the CD14 mutant mice did become insulin resistant and gained weight. Furthermore, hepatic steatosis, as reflected by the triglyceride content, was totally prevented in CD14 mutants during long-term high-fat feeding. Hepatic steatosis is an early event because only 4 weeks of high-fat feeding were sufficient to increase the triglyceride content of wild-type mice. This could be mimicked by a 4-week LPS infusion. Again, hepatic steatosis was totally blunted in CD14 mutant fed a high-fat diet or infused with LPS. We could conclude that the LPS-CD14 system sets a threshold at which metabolic diseases occur. Moreover, and relevant to our findings, epidemiological data from the literature showed statistical relationships between CD14 (39,40) inflammation, obesity, and insulin resistance in humans.

    LPS-treated mice developed inflammation, as the expression of genes coding for cytokines, IL-6, TNF-α, IL-1, and PAI-1 were increased in adipose depots, liver, and muscle. Importantly, these features occurred similarly in high-fat diet–fed mice. Interestingly, liver cytokines were increased as early as after 2 weeks of high-fat feeding, suggesting that the liver was most likely the first organ affected by LPS treatment. Evidence from the literature supports our hypothesis. It has been shown that LPS is a strong inducer of nonalcoholic hepatic steatosis (41), another feature related to metabolic diseases. In addition, genetically obese fa/fa rats and ob/ob mice quickly develop steatohepatitis after exposure to low doses of LPS (41). This is in agreement with a two- to four-times–increased endotoxemia characterizing db/db and ob/ob mice due to an increased intestinal permeability (42). Furthermore, polymyxin B treatment, which specifically eliminates Gram-negative bacterias and further quenches LPS, diminishes hepatic steatosis (43). Our data demonstrate that in absence of CD14, a high-fat diet could not induce inflammation as reflected by the total blunting of cytokine expression augmentation in the liver and adipose depot. Similarly, we recently reported that TLR4 mutant mice are protected against adipose tissue insulin resistance (44). Our finding is complementary to recent data that showed that free fatty acids can bind TLR4 to activate cells from the innate immune system (6) leading to the secretion of cytokines. This was totally blunted in TLR4 mutant mice. Therefore, our data suggest that TLR4 requires CD14, as a rate-limiting step, to mediate lipid-induced cytokine secretion because in the absence of CD14 high-fat diet could not induce inflammation in vivo; therefore, TLR4 is not sufficient. Another difference between the TLR4 and CD14 mutant mice is that the former became obese and hyperphagic, whereas CD14 mice delay the occurrence of obesity and showed no change in feeding behavior.

    Furthermore, whereas macrophage infiltration was increased in adipose tissue of wild-type mice fed a high-fat diet and in mice infused with LPS, this effect was totally blunted in absence of CD14. The mechanism involved in the recruitment of macrophages in the adipose depot seems to be different from the LPS coreceptor TLR4, as the corresponding mutant mice were still characterized by a high adipose tissue macrophage infiltration in response to high-fat diet (7,45). Conversely, TLR4 overexpression led to some extent of adipose insulin resistance (8). An original observation from our present work was that LPS increased the number of subcutaneous adipocytes of a smaller size. This is opposite to what observed in high-fat diet–fed mice. Again, CD14 mutant mice resist, to some extent, the changes in adipocyte size and number in response to high fat and LPS.

    The origin of metabolic diseases due to high-fat–induced metabolic endotoxemia in unclear but could be related to the microbiota present in the digestive tract. Recently, an original observation reported that young adult mice have 40% more total body fat than their germ-free counterpart fed the same diet (10,46). Similarly, lean axenic mice colonized with microbiota from ob/ob mice increased body weight. The authors suggested that gut microbiota from obese mice allows energy to be salvaged from otherwise-indigestible dietary polysaccharides. This hypothesis was further demonstrated by the fact that colonization increases glucose uptake in the host intestine and produces substantial elevations in serum glucose and insulin, both factors being lipogenic. However, this hypothesis of microbiota-induced lipogenesis could not explain either the inflammatory dimension of the mechanism linking dietary excess to obesity/diabetes or the differential effect on body weight of a high-fat diet versus a regular diet. However, axenic mice fed a high-fat obesitogenic diet did not gain weight, suggesting that, indeed, a bacterially related factor is responsible for high-fat diet–induced obesity (11). We suggest that LPS would be such a permissive factor that, upon binding to CD14, could lead to obesity. Therefore, lipids alone are not sufficient to promote obesity. Another recent report (12) showed that antibiotic treatment protected against the occurrence of autoimmune diabetes. The authors suggest that the gut microbiota generates inflammation and makes the rats prone to becoming diabetic. Hence, our data strongly support that metabolic endotoxemia would mediate the well-characterized low-tone inflammatory status of the metabolic disease. We suggest that LPS would be a newly identified inflammatory factor from microbiota, which upon binding to CD14 serves as vector for the triggering of obesity/insulin resistance induced by high-fat feeding. Along the same line of investigation, we previously reported that reducing the Gram-negative bacterial content of the digestive track by the mean of dietary fibers was also associated with reduced body weight (17,47).

    In conclusion, we demonstrate first that metabolic concentrations of plasma LPS are modulated by fat food content. Second, we found that metabolic concentrations of plasma LPS are a sufficient molecular mechanism for triggering the high-fat diet–induced metabolic diseases obesity/diabetes. Finally, the LPS receptor CD14, by controlling insulin sensitivity, sets the threshold at which metabolic diseases occur.

    FIG. 1.
    FIG. 1.

    High-fat feeding increased endotoxemia and changed intestinal microbiota. A: Plasma LPS concentration (EU/ml) was assessed every 4 h throughout the day in normal diet (CT; n = 9) (▪) and 4-week high-fat–fed (HF; n = 9) (○) mice. B: Groups of bacteria in the caecal content of mice fed the normal diet (CT; n = 8) or the high-fat diet (HF; n = 8) for 4 weeks. Bacterial numbers are expressed as log10 (bacterial cells per gram caecal content wet weight). *P < 0.05 vs. CT. C: Delta plasma LPS concentration in (EU/ml) in mice before and 30 min after an oral administration of LPS diluted in corn oil (n = 6) (oil-LPS) or in water (n = 6) (H2O-LPS) or an administration of oil alone (n = 6) (oil). *P < 0.05 vs. H2O-LPS. Data are means ± SE.

    FIG. 2.
    FIG. 2.

    Chronic experimental metabolic endotoxemia induces obesity and diabetes. A: Plasma endotoxin concentration (EU/ml) in WT mice infused with saline (CT; n = 18) or LPS (n = 18) for 4 weeks using subcutaneous osmotic pumps and compared with mice fed a high-fat diet for 4 weeks (HF; n = 18). B: Plasma glucose (mmol/l) following an oral glucose load (3 g/kg) in control (CT; n = 24), LPS (n = 13), or high-fat diet (HF; n = 24) mice. The inset represents the area under curve for each group. *P < 0.05 vs. CT; §LPS vs. CT; #HF vs. LPS. C: Plasma insulin (pmol/l) concentrations 30 min before (−30) and 15 min after (15) an oral glucose load in control (CT; n = 24), LPS (n = 13), or high-fat diet–fed (HF; n = 24) mice. D: Hepatic glucose production and whole-body glucose turnover rates (mg · kg−1 · min−1) in control (CT; n = 5), LPS (n = 5), or high-fat diet–fed (HF; n = 5) mice. E: Body weight (g) before (day 0) and after a 28-day treatment period (day 28) and body weight gain (Δ) in control (CT; n = 26), LPS (n = 21), or high-fat diet–fed (HF; n = 34) mice. F: Visceral and subcutaneous adipose tissue weight (percentage of body weight) in control (CT; n = 26), LPS (n = 21), or high-fat diet–fed (HF; n = 34) mice. G: Liver weight (percentage of body weight) in control (CT; n = 26), LPS (n = 21), or high-fat diet–fed (HF; n = 34) mice. H: Liver triglycerides (μmol/liver) in control (CT; n = 12), LPS (n = 9), or high-fat diet–fed (HF; n = 11) mice. I: Mean energy intake (kcal · day−1 · mouse−1) in control (CT; n = 18), LPS (n = 18), or high-fat diet–fed (HF; n = 18) mice. Data are means ± SE. Data with different superscript letters are significantly different at P < 0.05, according to the post hoc ANOVA statistical analysis.

    FIG. 3.
    FIG. 3.

    Metabolic endotoxemia triggers the expression of inflammatory factors similarly to high-fat feeding. TNF-α, IL-1, IL-6, and PAI-1 mRNA concentrations (A, E, and I) in liver (B, F, and J), visceral adipose tissue (C, G, and K), subcutaneous adipose tissue (D, H, and L), and muscle in normal diet–fed (n = 8) (□) or high-fat diet–fed (n = 8) (▪) mice for 2 weeks (AD) and 4 weeks (n = 8) (EH) and in LPS-infused mice (n = 5) (IL). Data are means ± SE. *P < 0.05 vs. normal chow–fed mice.

    FIG. 4.
    FIG. 4.

    CD14 mutant mice are protected against LPS-induced inflammation. mRNA concentrations of IL-6, PAI-1, and IL-1 in adipose tissue 3 h after a saline (control [CT]; n = 6) or an LPS (n = 6) infusion in WT (A) and CD14 mutant (B) mice. *P < 0.05 vs. CT; §P < 0.05 vs. WT. C: Representative Western blot analysis of p-NFk-B and p-IKK-b and p-IKK-a in the liver of mice from the same experiment. Protein Ct corresponds to a loading control of major protein, which cross-reacts nonspecifically with the anti–p-IKK-a antibody.

    FIG. 5.
    FIG. 5.

    The CD14 null mutation prevents the effect of LPS-induced obesity and diabetes. A: Body weight gain (g) in WT mice infused with saline (WT-CT; n = 13) or LPS (WT-LPS; n = 14) and CD14 mutant mice infused with saline (CD14-CT; n = 13) or LPS (CD14-LPS; n = 12) for 4 weeks using subcutaneous osmotic pumps. B: Visceral and subcutaneous adipose tissue weight (percentage of body weight) in WT-CT (n = 13) (□), WT-LPS (n = 14) (▪), CD14-CT (n = 13) (▒), and CD14-LPS (n = 12) (Embedded Image) mice. C: Plasma glucose concentration (mmol/l) following an intraperitoneal glucose load (1 g/kg) in WT-CT (n = 6) (▪), WT-LPS (n = 6) (▴), CD14-CT (n = 5) (□), and CD14-LPS (n = 6) (•) mice. The inset represents the area under curve of the same groups. D: Plasma insulin (pmol/l) concentration 30 min before (−30) and 30 min after (30) intraperitoneal glucose administration in WT-CT (n = 6) (□), WT-LPS (n = 6) (▪), CD14-CT (n = 5) (Embedded Image), and CD14-LPS (n = 6) (▒) mice. E: Liver weight (percentage of body weight) in WT-CT (n = 13), WT-LPS (n = 13), CD14-CT (n = 12), and CD14-LPS (n = 13) mice. F: Liver triglycerides (μmol/liver) in WT-CT (n = 12), WT-LPS (n = 9), CD14-CT (n = 5), and CD14-LPS (n = 6) mice. Data are means ± SE. Data with different superscript letters are significantly different at P < 0.05, according to the post hoc ANOVA statistical analysis.

    FIG. 6.

    CD14: the main LPS receptor sets the tone of insulin sensitivity and hepatic steatosis. A: Glucose turnover (mg · kg−1 · min−1) in WT mice or CD14 mutant fed a normal diet (CT) or fed a high-fat diet (HF) for 4 (n = 8) or 24 (n = 8) weeks (wks). B: Body weight gain (g) in WT or CD14 mutant fed a normal diet (CT) for 4 (n = 8) or 24 (n = 8) weeks or fed a high-fat diet (HF) for 4 (n = 8) or 24 (n = 8) weeks. C: Liver weight (percent of body weight) in WT or CD14 mutant mice fed a normal diet (CT) for 4 (n = 8) or 24 (n = 8) weeks or fed a high-fat diet (HF) for 4 (n = 8) or 24 (n = 8) weeks. D: Liver triglycerides (μmol/liver) in WT or CD14 mutant mice fed a normal diet (CT) for 4 (n = 6) or 24 (n = 5) weeks or fed a high-fat diet (HF) for 4 (n = 5) or 24 (n = 5) weeks. Data are means ± SE. *P < 0.05 vs. CT; §P < 0.05 vs. WT. E: Representative oil red O liver staining in 4-week–treated mice. F (see next page): Plasma glucose following an intraperitoneal glucose load (1 g/kg) in WT mice fed a normal diet (WT-CT; n = 8) or a high-fat diet (WT-HF; n = 8) for 24 weeks and CD14 mutant mice fed a normal diet (CD14-CT; n = 8) or fed a high-fat diet (CD14-HF; n = 8). The inset represents the area under curve of the same groups. G (see next page): Fasting plasma insulin (pmol/l) in WT or CD14 mutant mice fed a normal diet (CT) for 4 (n = 6) or 24 (n = 5) weeks or fed a high-fat diet (HF) for 4 (n = 5) or 24 (n = 5) weeks. Data are means ± SE. *P < 0.05 vs. CT; §P < 0.05 vs. WT. Data with different superscript letters are significantly different at P < 0.05, according to the post hoc ANOVA statistical analysis.

    FIG. 7.
    FIG. 7.

    CD14 mutant mice resist high-fat diet–induced glucose intolerance, inflammation, and increased visceral fat mass. A: Plasma glucose (mmol/l) following an intraperitoneal glucose load (1 g/kg) in WT mice fed a normal diet (WT-CT; n = 6) (▪) or a high-fat diet (WT-HF; n = 6) (▴) for 4 weeks and CD14 mutant mice fed a normal diet (CD14-CT; n = 5) (•) or fed a high-fat diet (CD14-HF; n = 5) (□). The inset represents the area under curve of the same groups. B: Plasma insulin concentration (pmol/l) 30 min before (−30) and 30 min after (30) intraperitoneal glucose administration in WT-CT (n = 6) (□), WT-HF (n = 6) (▪), CD14-CT (n = 5) ( ▒), and CD14-HF (n = 5) (Embedded Image) mice. C: Visceral and subcutaneous adipose tissue weight (percent of body weight) in WT-CT (n = 6) (□), WT-HF (n = 6) (▪), CD14-CT (n = 5) (▒), and CD14-HF (n = 5) (Embedded Image) mice. Adipose tissue (D) and liver (E) mRNA concentrations of TNF-α, IL-1, IL-6, and PAI-1 in WT-CT (n = 6) (□), WT-HF (n = 6) (▪), CD14-CT (n = 5) (▒), and CD14-HF (n = 5) (Embedded Image) mice. Data are means ± SE. Data with different superscript letters are significantly different at P < 0.05, according to the post hoc ANOVA statistical analysis.

    FIG. 8.

    Metabolic endotoxemia impairs adipocyte morphology. A: Adipocyte size distribution (%) in WT mice fed a normal diet (WT-CT; n = 765) (□) or a high-fat diet (WT-HF; n = 661) (▪) for 4 weeks or CD14 mutant mice fed a normal diet (CD14-CT; n = 1,032) ( ▒) or a high-fat diet (CD14-HF; n = 729) (Embedded Image) for 4 weeks. B: Adipocyte size distribution (%) in WT mice infused with saline (WT-CT; n = 765) (□) or LPS (WT-LPS; n = 750) (▪) and CD14 mutant mice infused with saline (CD14-CT; n = 1,032) (▒) or LPS (CD14-LPS; n = 585) (Embedded Image) for 4 weeks using subcutaneous osmotic pumps. C: Adipocyte mean area (μm2) in WT-CT (n = 765), WT-HF (n = 661), WT-LPS (n = 750), CD14-CT (n = 1,032), CD14-HF (n = 729), or CD14-LPS (n = 585) mice. D: Representative adipose tissue staining in 4-week–treated mice. E: F4/80-positive cells (+ cells/total cells) of all above groups. Data are means ± SE. Data with different superscript letters are significantly different at P < 0.05, according to the post hoc ANOVA statistical analysis.


    P.D.C. is a Postdoctoral Researcher from the FNRS (Fonds de la Recherche Scientifique), Belgium. R.B. is the recipient of subsides from the Nutritia Foundation, the AFERO (Association Française Etude et de Recherche sur les Obésités), the FRM (Fondation pour la Recherche Médicale), the ATIP (Action Thématiques Incitatives sur Programme), and the ACI (Action Concertée Incitative), Centre de la Recherche Nationale Scientifique, Université Paul Sabatier. J.A. is a recipient of a grant from the SFHTA (Société Française d’Hypertension Artérielle) and the délégation régionale à la recherche clinique des hôpitaux de Toulouse (in 2003). This work also was supported by the Programme National de Recherche sur les Maladies Cardiovasculaires, INSERM: “Vascular Risk and Metabolic Syndrome: Hemostasis, Inflammation and Metabolism Interplay” (no. A04046AS) (to M.C.A.). This work was supported by a Fonds Speciaux de Recherche grant from the Université catholique de Louvain and FNRS grant to N.M.D.

    We thank Drs. B. Pipy, J.F. Arnal, C. Feyt, and P. Gourdy for helpful criticisms and N. Maton and C. Dray for excellent technical assistance.


    • Published ahead of print at on 24 April 2007. DOI: 10.2337/db06-1491.

    • P.D.C., J.A., and M.A.I. contributed equally to this article.

    • The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

      • Accepted April 13, 2007.
      • Received October 24, 2006.


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    Sell Unused Diabetic Strips Today!

    Easy Homemade Low-Sodium Salsa – Diabetic Foodie

    By electricdiet / August 10, 2021

    This easy homemade low-sodium salsa is packed with all the fresh ingredients and vibrant flavors you love without the extra sodium or fillers you find in the store-bought versions!

    Bowl of homemade low-sodium salsa on bed of nacho chips

    When you start cutting salt out of your diet, you might be surprised to see how much sodium is in certain foods.

    Some of the biggest culprits are processed foods like salty snacks, canned soup, or cured meats. But I was surprised to find that two tablespoons of Tostitos Chunky Salsa – Medium has 250mg of sodium!

    Many brands have added sugar and fillers as well.

    That’s why I love making this easy low-sodium salsa right at home! By comparison, it only has 4mg of sodium per two tablespoons. And of course, there’s no added sugar.

    Salsa is so delicious to enjoy as a dip or in recipes like Nacho Celery Sticks, or Mexican Microwave Egg Scramble. Plus, it whips up in just minutes. I can have salsa ready to go any time!

    If you’ve been watching your sodium intake or just want an easy, fresh salsa without unnecessary fillers or sugar, you’ll love this recipe.

    How to make low-sodium salsa

    Homemade salsa is so delicious and simple to make, you’ll never want to buy it from the store ever again! Plus, this recipe has way less sodium, contains no artificial ingredients, and is very affordable!

    Salsa ingredients on a cutting board

    Step 1: Add the cleaned and roughly chopped tomatoes, onions, jalapeño, cilantro, and lime juice to your food processor.

    Ingredients in a food processor

    Step 2: Pulse until the salsa reaches your desired consistency. I like it to still be a little bit chunky.

    Step 3: Garnish with cilantro if desired and serve!

    Chip with salsa over a bowl of salsa

    Playing with the flavor of your salsa

    One of my favorite things about salsa is how many different variations you can make. You can customize it to your tastes and have some fun!

    If you prefer a less spicy salsa, swap out the jalapeño for a milder pepper or bell pepper. On the other hand, if you love spice, you could use something hotter like serrano peppers instead!

    You can also use a variety of onions. I really like green onions, so that’s what I use here, but red onions are also very popular in salsa recipes. Feel free to use whatever you have on hand.

    When picking out tomatoes at the store or farmers’ market, you can choose whichever variety looks the best! Tomatoes make up the bulk of salsa, so the better the tomato tastes, the better your salsa will be. If you can find them, I especially love heirloom tomatoes.

    And finally, if you belong to the part of the population that thinks cilantro tastes like soap, you’re welcome to swap it out for parsley instead!

    Getting the right texture

    Just like the flavors, you can also customize the texture of the salsa to your preference.

    If you like it super smooth, simply pulse for longer in the food processor. On the other hand, if you prefer really chunky salsa, you can skip the food processor and chop everything by hand!

    Bowl of low-sodium salsa with sliced jalapenos on top


    Your homemade salsa can be stored in an airtight container in the refrigerator. For the best flavor, I recommend eating it within 3 days.

    The cilantro will wilt the fastest, so if you leave it out, you could store the salsa for up to 5 days in the refrigerator and simply add the cilantro right before serving.

    More healthy recipes to try

    Looking for more delicious salsa recipes or dip recipes to spice up Taco Tuesday? Here are a few more dishes I know you’ll love:

    When you’ve tried this salsa please don’t forget to let me know how you liked it and rate the recipe in the comments below!

    Recipe Card

    Low-sodium Easy Homemade Salsa

    Low-Sodium Easy Homemade Salsa

    Easy-to-make salsa with much less sodium than store-bought

    Prep Time:10 minutes

    Total Time:10 minutes

    Author:Diabetic Foodie



    • Add the tomatoes, onions, jalapeño, cilantro, and lime juice to your food processor.

    • Pulse until the salsa reaches your desired consistency.

    • Garnish with cilantro if desired and serve!

    Recipe Notes

    This recipe is for 8 servings of salsa. Each serving will be about 2 tablespoons.
    1 tablespoon of yellow or red onion can be used in place of the green onions.
    For a milder salsa, replace the jalapeño with about 1 tablespoon banana pepper or green bell pepper. For a hotter salsa, use a spicier pepper like a serrano.
    For a really chunky salsa, dice everything by hand and skip the food processor.
    Salsa can be stored in an airtight container in the refrigerator. Consume within 3 days for the best flavor.

    Nutrition Info Per Serving

    Nutrition Facts

    Low-Sodium Easy Homemade Salsa

    Amount Per Serving (2 tablespoons)

    Calories 10

    % Daily Value*

    Fat 0g0%

    Saturated Fat 0g0%

    Trans Fat 0g

    Polyunsaturated Fat 0g

    Monounsaturated Fat 0g

    Cholesterol 0mg0%

    Sodium 6.8mg0%

    Potassium 65.3mg2%

    Carbohydrates 2g1%

    Fiber 0.3g1%

    Sugar 0.9g1%

    Protein 0.3g1%

    Vitamin A 550IU11%

    Vitamin C 9.9mg12%

    Calcium 10mg1%

    Iron 0.2mg1%

    Net carbs 1.7g

    * Percent Daily Values are based on a 2000 calorie diet.

    Course: Appetizer

    Cuisine: Mexican

    Diet: Diabetic, Gluten Free

    Keyword: homemade salsa, low sodium, low-sodium salsa

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